Cell and Molecular Biology Concepts and Experiments 7th Edition Karp Solutions Manual

1. 1: Introduction to the Study of Cell and Molecular Biology
2. An example of the role of technological innovation in the field of cell biology. This light micrograph shows a cell lying on a microscopic bed of synthetic posts. The flexible posts serve as sensors to measure mechanical forces exerted by the cell. The red-stained elements are bundles of actin filaments within the cell that generate forces during cell locomotion. When the cell moves, it pulls on the attached posts, which report the amount of strain they are experiencing. The cell nucleus is stained green.
3. 1.1: The Discovery of Cells
4. Figure 1.1: The discovery of cells. (a) One of Robert Hooke’s more ornate compound (double-lens) microscopes. (Inset) Hooke’s drawing of a thin slice of cork, showing the honeycomb-like network of “cells.” (b) Single-lens microscope used by Anton van Leeuwenhoek to observe bacteria and other microorganisms. The biconvex lens, which was capable of magnifying an object approximately 270 times and providing a resolution of approximately 1.35 μm, was held between two metal plates.
5. 1.2: Basic Properties of Cells
6. Cells Are Highly Complex and Organized
7. Figure 1.2: HeLa cells, such as the ones pictured here, were the first human cells to be kept in culture for long periods of time and are still in use today. Unlike normal cells, which have a finite lifetime in culture, these cancerous HeLa cells can be cultured indefinitely as long as conditions are favorable to support cell growth and division.
8. Figure 1.3: Levels of cellular and molecular organization. The brightly colored photograph of a stained section shows the microscopic structure of a villus of the wall of the small intestine, as seen through the light microscope. Inset 1 shows an electron micrograph of the epithelial layer of cells that lines the inner intestinal wall. The apical surface of each cell, which faces the channel of the intestine, contains a large number of microvilli involved in nutrient absorption. The basal region of each cell contains large numbers of mitochondria, where energy is made available to the cell. Inset 2 shows the apical microvilli; each microvillus contains a bundle of microfilaments. Inset 3 shows the actin protein subunits that make up each microfilament. Inset 4 shows an individual mitochondrion similar to those found in the basal region of the epithelial cells. Inset 5 shows a portion of an inner membrane of a mitochondrion including the stalked particles (upper arrow) that project from the membrane and correspond to the sites where ATP is synthesized. Insets 6 and 7 show molecular models of the ATP-synthesizing machinery, which is discussed at length in Chapter 5.
9. Cells Possess a Genetic Program and the Means to Use It
10. Cells Are Capable of Producing More of Themselves
11. Figure 1.4: Cell reproduction. This mammalian oocyte has recently undergone a highly unequal cell division in which most of the cytoplasm has been retained within the large oocyte, which has the potential to be fertilized and develop into an embryo. The other cell is a nonfunctional remnant that consists almost totally of nuclear material (indicated by the blue-staining chromosomes, arrow).
12. Cells Acquire and Utilize Energy
13. Figure 1.5: Acquiring energy. A living cell of the filamentous alga Spirogyra. The ribbon-like chloroplast, which is seen to zigzag through the cell, is the site where energy from sunlight is captured and converted to chemical energy during photosynthesis.
14. Cells Carry Out a Variety of Chemical Reactions
15. Cells Engage in Mechanical Activities
16. Cells Are Able to Respond to Stimuli
17. Cells Are Capable of Self-Regulation
18. Figure 1.6: Self-regulation. The left panel depicts the normal development of a sea urchin in which a fertilized egg gives rise to a single embryo. The right panel depicts an experiment in which the cells of an early embryo are separated from one another after the first division, and each cell is allowed to develop in isolation. Rather than developing into half of an embryo, as it would if left undisturbed, each isolated cell recognizes the absence of its neighbor, regulating its development to form a complete (although smaller) embryo.
19. Figure 1.7: Cellular activities are often analogous to this “Rube Goldberg machine” in which one event “automatically” triggers the next event in a reaction sequence.
20. Cells Evolve
21. REVIEW
22. 1.3: Two Fundamentally Different Classes of Cells
23. Figure 1.8: The structure of cells. Schematic diagrams of a “generalized” bacterial (a), plant (b), and animal (c) cell. Note: Organelles are not drawn to scale.
24. Characteristics That Distinguish Prokaryotic and Eukaryotic Cells
25. Figure 1.9: Earth’s biogeologic clock. A portrait of the past five billion years of Earth’s history showing a proposed time of appearance of major groups of organisms. Complex animals (shelly invertebrates) and vascular plants are relatively recent arrivals. The time indicated for the origin of life is speculative. In addition, photosynthetic bacteria may have arisen much earlier, hence the question mark. The geologic eras are indicated in the center of the illustration.
26. Table 1.1: A Comparison of Prokaryotic and Eukaryotic Cells
27. Figure 1.10: The structure of a eukaryotic cell. This epithelial cell lines the male reproductive tract in the rat. A number of different organelles are indicated and depicted in schematic diagrams around the border of the figure.
28. Figure 1.11: The cytoplasm of a eukaryotic cell is a crowded compartment. This colorized electron micrographic image shows a small region near the edge of a single-celled eukaryotic organism that had been quickly frozen prior to microscopic examination. The three-dimensional appearance is made possible by capturing two-dimensional digital images of the specimen at different angles and merging the individual frames using a computer. Cytoskeletal filaments are shown in red, macromolecular complexes (primarily ribosomes) are green, and portions of cell membranes are blue.
29. Figure 1.12: Cell division in eukaryotes requires the assembly of an elaborate chromosome-separating apparatus called the mitotic spindle, which is constructed primarily of microtubules. The microtubules in this micrograph appear green because they are bound by an antibody that is linked to a green fluorescent dye. The chromosomes, which were about to be separated into two daughter cells when this cell was fixed, are stained blue.
30. Figure 1.13: Bacterial conjugation. Electron micrograph showing a conjugating pair of bacteria joined by a structure of the donor cell, termed the F pilus, through which DNA is thought to be passed.
31. Figure 1.14: The difference between prokaryotic and eukaryotic flagella.(a) The bacterium Salmonella with its numerous flagella. Inset shows a high-magnification view of a portion of a single bacterial flagellum, which consists largely of a single protein called flagellin. (b) Each of these human sperm cells is powered by the undulatory movements of a single flagellum. The inset shows a cross section of the central core of a mammalian sperm flagellum. The flagella of eukaryotic cells are so similar that this cross section could just as well have been taken of a flagellum from a protist or green alga.
32. Types of Prokaryotic Cells
33. Figure 1.15: Cyanobacteria.(a) Electron micrograph of a cyanobacterium showing the cytoplasmic membranes that carry out photosynthesis. These concentric membranes are very similar to the thylakoid membranes present within the chloroplasts of plant cells, a reminder that chloroplasts evolved from a symbiotic cyanobacterium. (b) Cyanobacteria living inside the hairs of these polar bears are responsible for the unusual greenish color of their coats.
34. Prokaryotic Diversity
35. Table 1.2: Number and Biomass of Prokaryotes in the World
36. Figure 1.16: Vorticella, a complex ciliated protist. A number of these unicellular organisms are seen here; most have withdrawn their “heads” due to shortening of the blue-stained contractile ribbon in the stalk. Each cell has a single large nucleus, called a macronucleus (arrow), which contains many copies of the genes.
37. Types of Eukaryotic Cells: Cell Specialization
38. Figure 1.17: Pathways of cell differentiation. A few of the types of differentiated cells present in a human fetus.
39. Model Organisms
40. The Sizes of Cells and Their Components
41. Synthetic Biology
42. Figure 1.18: Six model organisms. (a) Escherichia coli is a rod-shaped bacterium that lives in the digestive tract of humans and other mammals. Much of what we will discuss about the basic molecular biology of the cell, including the mechanisms of replication, transcription, and translation, was originally worked out on this one prokaryotic organism. The relatively simple organization of a prokaryotic cell is illustrated in this electron micrograph (compare to part b of a eukaryotic cell). (b) Saccharomyces cerevisiae, more commonly known as baker’s yeast or brewer’s yeast. It is the least complex of the eukaryotes commonly studied, yet it contains a surprising number of proteins that are homologous to proteins in human cells. Such proteins typically have a conserved function in the two organisms. The species has a small genome encoding about 6200 proteins; it can be grown in a haploid state (one copy of each gene per cell rather than two as in most eukaryotic cells); and it can be grown under either aerobic (O2-containing) or anaerobic (O2-lacking) conditions. It is ideal for the identification of genes through the use of mutants. (c) Arabidopsis thaliana, a weed (called the thale cress) that is related to mustard and cabbage, which has an unusually small genome (120 million base pairs) for a flowering plant, a rapid generation time, and large seed production, and it grows to a height of only a few inches. (d) Caenorhabditis elegans, a microscopic-sized nematode, consists of a defined number of cells (roughly 1000), each of which develops according to a precise pattern of cell divisions. The animal is easily cultured, can be kept alive in a frozen state, has a transparent body wall, a short generation time, and facility for genetic analysis. This micrograph shows the larval nervous system, which has been labeled with the green fluorescent protein (GFP). The 2002 Nobel Prize was awarded to the researchers who pioneered its study. (e) Drosophila melanogaster, the fruit fly, is a small but complex eukaryote that is readily cultured in the lab, where it grows from an egg to an adult in a matter of days. Drosophila has been a favored animal for the study of genetics, the molecular biology of development, and the neurological basis of simple behavior. Certain larval cells have giant chromosomes, whose individual genes can be identified for studies of evolution and gene expression. In the mutant fly shown here, a leg has developed where an antenna would be located in a normal (wild type) fly. (f) Mus musculus, the common house mouse, is easily kept and bred in the laboratory. Thousands of different genetic strains have been developed, many of which are stored simply as frozen embryos due to lack of space to house the adult animals. The “nude mouse” pictured here develops without a thymus gland and, therefore, is able to accept human tissue grafts that are not rejected.
43. Figure 1.19: Relative sizes of cells and cell components. These structures differ in size by more than seven orders of magnitude.
44. Figure 1.20: The synthetic biologist’s toolkit of the future? Such a toolkit would presumably contain nucleic acids, proteins, lipids, and many other types of biomolecules.
45. REVIEW
46. THE HUMAN PERSPECTIVE: The Prospect of Cell Replacement Therapy
47. Adult Stem Cells
48. Figure 1: An adult muscle stem cell. (a) A portion of a muscle fiber, with its many nuclei stained blue. A single stem cell (yellow) is seen to be lodged between the outer surface of the muscle fiber and an extracellular layer (or basement membrane), which is stained red. The undifferentiated stem cell exhibits this yellow color because it expresses a protein that is not present in the differentiated muscle fiber. (b) Adult stem cells undergoing differentiation into adipose (fat) cells in culture. Stem cells capable of this process are present in adult fat tissue and also bone marrow.
49. Embryonic Stem Cells
50. Figure 2: Embryonic stem cells; their isolation and potential use. (a) Micrograph of a mammalian blastocyst, an early stage during embryonic development, showing the inner cell mass, which is composed of pluripotent ES cells. Once isolated, such cells are readily grown in culture. (b) A potential procedure for obtaining differentiated cells for use in cell replacement therapy. A small piece of tissue is taken from the patient, and one of the somatic cells is fused with a donor oocyte whose own nucleus had been previously removed. The resulting oocyte (egg), with the patient’s cell nucleus, is allowed to develop into an early embryo, and the ES cells are harvested and grown in culture. A population of ES cells are induced to differentiate into the required cells, which are subsequently transplanted into the patient to restore organ function. (At the present time, it has not been possible to obtain blastocyst stage embryos, that is, ones with ES cells, from any primate species by the procedure shown here, although it has been accomplished using an oocyte from which the nucleus is not first removed. The ES cells that are generated in such experiments are triploid; that is, they have three copies of each chromosome—one from the oocyte and two from the donor nucleus—rather than two, as would normally be the case. Regardless, these triploid ES cells are pluripotent and capable of transplantation.)
51. Induced Pluripotent Stem Cells
52. Figure 3: Steps taken to generate induced pluripotent stem (iPS) cells for use in correcting the inherited disease sickle cell anemia in mice. Skin cells are collected from the diseased animal, reprogrammed in culture by introducing the four required genes that are ferried into the cells by viruses, and allowed to develop into undifferentiated pluripotent iPS cells. The iPS cells are then treated so as to replace the defective (globin) gene with a normal copy, and the corrected iPS cells are caused to differentiate into normal blood stem cells in culture. These blood stem cells are then injected back into the diseased mouse, where they proliferate and differentiate into normal blood cells, thereby curing the disorder.
53. Direct Cell Reprogramming
54. 1.4: Viruses
55. Figure 1.21: Tobacco mosaic virus (TMV). (a) Model of a portion of a TMV particle. The protein subunits, which are identical along the entire rod-shaped particle, enclose a single helical RNA molecule (red). (b) Electron micrograph of TMV particles after phenol has removed the protein subunits from the middle part of the upper particle and the ends of the lower particle. Intact rods are approximately 300 nm long and 18 nm in diameter.
56. Figure 1.22: Virus diversity. The structures of (a) an adenovirus, (b) a human immunodeficiency virus (HIV), and (c) a T-even bacteriophage.
57. Figure 1.23: A virus infection. (a) Micrograph showing a late stage in the infection of a bacterial cell by a bacteriophage. Virus particles are being assembled within the cell, and empty phage coats are still present on the cell surface. (b) Micrograph showing HIV particles budding from an infected human lymphocyte.
58. Viroids
59. REVIEW
60. EXPERIMENTAL PATHWAYS: The Origin of Eukaryotic Cells
61. Figure 1: A model depicting possible steps in the evolution of eukaryotic cells, including the origin of mitochondria and chloroplasts by endosymbiosis. In step 1, a large anaerobic, heterotrophic prokaryote takes in a small aerobic prokaryote. Evidence strongly indicates that the engulfed prokaryote was an ancestor of modern-day rickettsia, a group of bacteria that causes typhus and other diseases. In step 2, the aerobic endosymbiont has evolved into a mitochondrion. In step 3, a portion of the plasma membrane has invaginated and is seen in the process of evolving into a nuclear envelope and associated endoplasmic reticulum. The pre-eukaryote depicted in step 3 gives rise to two major groups of eukaryotes. In one path (step 4), a primitive eukaryote evolves into nonphotosynthetic protist, fungal, and animal cells. In the other path (step 5), a primitive eukaryote takes in a photosynthetic prokaryote, which will become an endosymbiont and evolve into a chloroplast.
62. Figure 2: Two-dimensional electrophoretic fingerprint of a T1 digest of chloroplast 16S ribosomal RNA. The RNA fragments were electrophoresed in one direction at pH 3.5 and then in a second direction at pH 2.3.
63. Table 1: Nucleotide Sequence Similarities between Representative Members of the Three Primary Kingdoms
64. Figure 3: A phylogenetic tree based on rRNA sequence comparisons showing the three domains of life. The Archaea are divided into two subgroups as indicated.
65. References
66. Synopsis
67. Analytic Questions
68. 2: The Chemical Basis of Life
69. A complex between two different macromolecules. A portion of a DNA molecule (shown in blue) is complexed to a protein that consists of two polypeptide subunits, one red and the other yellow. Those parts of the protein that are seen to be inserted into the grooves of the DNA have recognized and bound to a specific sequence of nucleotides in the nucleic acid molecule.
70. 2.1: Covalent Bonds
71. Figure 2.1: A representation of the arrangement of electrons in a number of common atoms. Electrons are present around an atom’s nucleus in “clouds” or orbitals that are roughly defined by their boundaries, which may have a spherical or dumbbell shape. Each orbital contains a maximum of two electrons, which is why the electrons (dark dots in the drawing) are grouped in pairs. The innermost shell contains a single orbital (thus two electrons), the second shell contains four orbitals (thus eight electrons), the third shell also contains four orbitals, and so forth. The number of outer-shell electrons is a primary determinant of the chemical properties of an element. Atoms with a similar number of outer-shell electrons have similar properties. Lithium (Li) and sodium (Na), for example, have one outer-shell electron, and both are highly reactive metals. Carbon (C) and silicon (Si) atoms can each bond with four different atoms. Because of its size, however, a carbon atom can bond to other carbon atoms, forming long-chained organic molecules, whereas silicon is unable to form comparable molecules. Neon (Ne) and argon (Ar) have filled outer shells, making these atoms highly nonreactive; they are referred to as inert gases.
72. Polar and Nonpolar Molecules
73. Ionization
74. REVIEW
75. 2.2: Noncovalent Bonds
76. THE HUMAN PERSPECTIVE: Free Radicals as a Cause of Aging
77. Ionic Bonds: Attractions between Charged Atoms
78. Figure 2.2: The dissolution of a salt crystal. When placed in water, the Na+ and Cl− ions of a salt crystal become surrounded by water molecules, breaking the ionic bonds between the two ions. As the salt dissolves, the negatively charged oxygen atoms of the water molecules associate with the positively charged sodium ions, and the positively charged hydrogen atoms of the water molecules associate with the negatively charged chloride ions.
79. Hydrogen Bonds
80. Figure 2.3: Noncovalent ionic bonds play an important role in holding the protein molecule on the right (yellow atoms) to the DNA molecule on the left. Ionic bonds form between positively charged nitrogen atoms in the protein and negatively charged oxygen atoms in the DNA. The DNA molecule itself consists of two separate strands held together by noncovalent hydrogen bonds. Although a single noncovalent bond is relatively weak and easily broken, large numbers of these bonds between two molecules, as between two strands of DNA, make the overall complex quite stable.
81. Hydrophobic Interactions and van der Waals Forces
82. Figure 2.4: Hydrogen bonds form between a bonded electronegative atom, such as nitrogen or oxygen, which bears a partial negative charge, and a bonded hydrogen atom, which bears a partial positive charge. Hydrogen bonds (about 0.18 nm) are typically about twice as long as the much stronger covalent bonds.
83. Figure 2.5: In a hydrophobic interaction, the nonpolar (hydrophobic) molecules are forced into aggregates, which minimizes their exposure to the surrounding water molecules.
84. The Life-Supporting Properties of Water
85. Figure 2.6: Van der Waals forces.(a) As two atoms approach each other, they experience a weak attractive force that increases up to a specific distance, typically about 4 Å. If the atoms approach more closely, their electron clouds repel one another, causing the atoms to be forced apart. (b) Although individual van der Waals forces are very weak and transient, large numbers of such attractive forces can be formed if two macromolecules have a complementary surface, as is indicated schematically in this figure (see Figure 2.40 for an example).
86. Figure 2.7: Hydrogen bond formation between neighboring water molecules. Each H atom of the molecule has about four-tenths of a full positive charge, and the single O atom has about eight-tenths of a full negative charge.
87. REVIEW
88. Figure 2.8: The importance of water in protein structure. The water molecules (each with a single red oxygen atom and two smaller gray hydrogen atoms) are shown in their ordered locations between the two subunits of a clam hemoglobin molecule.
89. 2.3: Acids, Bases, and Buffers
90. Table 2.1: Strengths of Acids and Bases
91. REVIEW
92. 2.4: The Nature of Biological Molecules
93. Figure 2.9: Cholesterol, whose structure illustrates how carbon atoms (represented by the black balls) are able to form covalent bonds with as many as four other carbon atoms. As a result, carbon atoms can be linked together to form the backbones of a virtually unlimited variety of organic molecules. The carbon backbone of a cholesterol molecule includes four rings, which is characteristic of steroids (e.g., estrogen, testosterone, cortisol). The cholesterol molecule shown here is drawn as a ball-and-stick model, which is another way that molecular structure is depicted.
94. Functional Groups
95. Table 2.2: Functional Groups
96. A Classification of Biological Molecules by Function
97. Figure 2.10: Monomers and polymers; polymerization and hydrolysis. a) Polysaccharides, proteins, and nucleic acids consist of monomers (subunits) linked together by covalent bonds. Free monomers do not simply react with each other to become macromolecules. Rather, each monomer is first activated by attachment to a carrier molecule that subsequently transfers the monomer to the end of the growing macromolecule.(b) A macromolecule is disassembled by hydrolysis of the bonds that join the monomers together. Hydrolysis is the splitting of a bond by water. All of these reactions are catalyzed by specific enzymes.
98. REVIEW
99. 2.5: Four Types of Biological Molecules
100. Figure 2.11: An overview of the types of biological molecules that make up various cellular structures.
101. Carbohydrates
102. The Structure of Simple Sugars
103. Figure 2.12: The structures of sugars.(a) Straight-chain formula of fructose, a ketohexose [keto, indicating the carbonyl (yellow), is located internally, and hexose because it consists of six carbons]. (b) Straight-chain formula of glucose, an aldohexose (aldo because the carbonyl is located at the end of the molecule). (c) Self-reaction in which glucose is converted from an open chain to a closed ring (a pyranose ring). (d) Glucose is commonly depicted in the form of a flat (planar) ring lying perpendicular to the page with the thickened line situated closest to the reader and the H and OH groups projecting either above or below the ring. The basis for the designation α-D-glucose is discussed in the following section. (e) The chair conformation of glucose, which depicts its three-dimensional structure more accurately than the flattened ring of part d. (f) A ball-and-stick model of the chair conformation of glucose.
104. Stereoisomerism
105. Figure 2.13: Stereoisomerism of glyceraldehyde.(a) The four groups bonded to a carbon atom (labeled a, b, c, and d) occupy the four corners of a tetrahedron with the carbon atom at its center. (b) Glyceraldehyde is the only three-carbon aldose; its second carbon atom is bonded to four different groups (—H, —OH, —CHO, and —CH2OH). As a result, glyceraldehyde can exist in two possible configurations that are not superimposable, but instead are mirror images of each other as indicated. These two stereoisomers (or enantiomers) can be distinguished by the configuration of the four groups around the asymmetric (or chiral) carbon atom. Solutions of these two isomers rotate plane-polarized light in opposite directions and, thus, are said to be optically active. (c) Straight-chain formulas of glyceraldehyde. By convention, the D-isomer is shown with the OH group on the right.
106. Figure 2.14: Aldotetroses. Because they have two asymmetric carbon atoms, aldotetroses can exist in four configurations.
107. Linking Sugars Together
108. Figure 2.15: Formation of an α- and β-pyranose. When a molecule of glucose undergoes self-reaction to form a pyranose ring (i.e., a six-membered ring), two stereoisomers are generated. The two isomers are in equilibrium with each other through the open-chain form of the molecule. By convention, the molecule is an α-pyranose when the OH group of the first carbon projects below the plane of the ring, and a β-pyranose when the hydroxyl group projects upward.
109. Figure 2.16: Disaccharides. Sucrose and lactose are two of the most common disaccharides. Sucrose is composed of glucose and fructose joined by an α(1→2) linkage, whereas lactose is composed of glucose and galactose joined by a β(1→4) linkage.
110. Polysaccharides
111. Glycogen and Starch: Nutritional Polysaccharides
112. Figure 2.17: Three polysaccharides with identical sugar monomers but dramatically different properties. Glycogen (a), starch (b), and cellulose (c) are each composed entirely of glucose subunits, yet their chemical and physical properties are very different due to the distinct ways that the monomers are linked together (three different types of linkages are indicated by the circled numbers). Glycogen molecules are the most highly branched, starch molecules assume a helical arrangement, and cellulose molecules are unbranched and highly extended. Whereas glycogen and starch are energy stores, cellulose molecules are bundled together into tough fibers that are suited for their structural role. Colorized electron micrographs show glycogen granules in a liver cell, starch grains (amyloplasts) in a plant seed, and cellulose fibers in a plant cell wall; each is indicated by an arrow.
113. Cellulose, Chitin, and Glycosaminoglycans: Structural Polysaccharides
114. Figure 2.18: Chitin is the primary component of the outer skeleton of this grasshopper.
115. Lipids
116. Fats
117. Figure 2.19: Fats and fatty acids.(a) The basic structure of a triacylglycerol (also called a triglyceride or a neutral fat). The glycerol moiety, indicated in orange, is linked by three ester bonds to the carboxyl groups of three fatty acids whose tails are indicated in green. (b) Stearic acid, an 18-carbon saturated fatty acid that is common in animal fats. (c) Space-filling model of tristearate, a triacylglycerol containing three identical stearic acid chains. (d) Space-filling model of linseed oil, a triacylglycerol derived from flax seeds that contains three unsaturated fatty acids (linoleic, oleic, and linolenic acids). The sites of unsaturation, which produce kinks in the molecule, are indicated by the yellow-orange bars.
118. Figure 2.20: Soaps consist of fatty acids. In this schematic drawing of a soap micelle, the nonpolar tails of the fatty acids are directed inward, where they interact with the greasy matter to be dissolved. The negatively charged heads are located at the surface of the micelle, where they interact with the surrounding water. Membrane proteins, which also tend to be insoluble in water, can also be solubilized in this way by extraction of membranes with detergents.
119. Figure 2.21: The structure of steroids. All steroids share the basic four-ring skeleton. The seemingly minor differences in chemical structure between cholesterol, testosterone, and estrogen generate profound biological differences.
120. Steroids
121. Phospholipids
122. Figure 2.22: The phospholipid phosphatidylcholine. The molecule consists of a glycerol backbone whose hydroxyl groups are covalently bonded to two fatty acids and a phosphate group. The negatively charged phosphate is also bonded to a small, positively charged choline group. The end of the molecule that contains the phosphorylcholine is hydrophilic, whereas the opposite end, consisting of the fatty acid tail, is hydrophobic. The structure and function of phosphatidylcholine and other phospholipids are discussed at length in Section 4.3.
123. Proteins
124. The Building Blocks of Proteins
125. Figure 2.23: Two examples of the thousands of biological structures composed predominantly of protein. These include (a) feathers, which are adaptations in birds for thermal insulation, flight, and sex recognition; and (b) the lenses of eyes, as in this spider, which focus light rays.
126. The Structures of Amino Acids
127. Figure 2.24: Amino acid structure. Ball-and-stick model (a) and chemical formula (b) of a generalized amino acid in which R can be any of a number of chemical groups (see Figure 2.26).(c) The formation of a peptide bond occurs by the condensation of two amino acids, drawn here in the uncharged state. In the cell, this reaction occurs on a ribosome as an amino acid is transferred from a carrier (a tRNA molecule) onto the end of the growing polypeptide chain (see Figure 11.49).
128. Figure 2.25: Amino acid stereoisomerism. Because the α-carbon of all amino acids except glycine is bonded to four different groups, two stereoisomers can exist. The D and L forms of alanine are shown.
129. The Properties of the Side Chains
130. Figure 2.26: The chemical structure of amino acids. These 20 amino acids represent those most commonly found in proteins and, more specifically, those encoded by DNA. Other amino acids occur as the result of a modification to one of those shown here. The amino acids are arranged into four groups based on the character of their side chains, as described in the text. All molecules are depicted as free amino acids in their ionized state as they would exist in solution at neutral pH.
131. Figure 2.27: The ionization of charged, polar amino acids.(a) The side chain of glutamic acid loses a proton when its carboxylic acid group ionizes. The degree of ionization of the carboxyl group depends on the pH of the medium: the greater the hydrogen ion concentration (the lower the pH), the smaller the percentage of carboxyl groups that are present in the ionized state. Conversely, a rise in pH leads to an increased ionization of the proton from the carboxyl group, increasing the percentage of negatively charged glutamic acid side chains. The pH at which 50 percent of the side chains are ionized and 50 percent are unionized is called the pK, which is 4.4 for the side chain of free glutamic acid. At physiologic pH, virtually all of the glutamic acid residues of a polypeptide are negatively charged. (b) The side chain of lysine becomes ionized when its amino group gains a proton. The greater the hydroxyl ion concentration (the higher the pH), the smaller the percentage of amino groups that are positively charged. The pH at which 50 percent of the side chains of lysine are charged and 50 percent are uncharged is 10.0, which is the pK for the side chain of free lysine. At physiologic pH, virtually all of the lysine residues of a polypeptide are positively charged. Once incorporated into a polypeptide, the pK of a charged group can be greatly influenced by the surrounding environment.
132. Figure 2.28: Disposition of hydrophilic and hydrophobic amino acid residues in the soluble protein cytochrome c. (a) The hydrophilic side chains, which are shown in green, are located primarily at the surface of the protein where they contact the surrounding aqueous medium. (b) The hydrophobic residues, which are shown in red, are located primarily within the center of the protein, particularly in the vicinity of the central heme group.
133. The Structure of Proteins
134. Figure 2.29: Scanning electron micrograph of a red blood cell from a person with sickle cell anemia. Compare with the micrograph of a normal red blood cell of Figure 2.32a.
135. Primary Structure
136. Secondary Structure
137. Figure 2.30: The alpha helix. (a) Tubular representation of an α helix. The atoms of the main chain would just fit within a tube of this radius. (b) The helical path around a central axis taken by the polypeptide backbone in a region of α helix. Each complete (360°) turn of the helix corresponds to 3.6 amino acid residues. The distance along the axis between adjacent residues is 1.5A. (c) The arrangement of the atoms of the backbone of the α helix and the hydrogen bonds that form between amino acids. Because of the helical rotation, the peptide bonds of every fourth amino acid come into close proximity. The approach of the carbonyl group (C=O) of one peptide bond to the imine group (H—N) of another peptide bond results in the formation of hydrogen bonds between them. The hydrogen bonds (orange bars) are essentially parallel to the axis of the cylinder and thus hold the turns of the chain together.
138. Figure 2.31: The β-pleated sheet. (a) Tubular representation of an antiparallel β sheet. (b) Each polypeptide of a β sheet assumes an extended but pleated conformation referred to as a β strand. The pleats result from the location of the α-carbons above and below the plane of the sheet. Successive side chains (R groups in the figure) project upward and downward from the backbone. The distance along the axis between adjacent residues is 3.5Å (c) A β-pleated sheet consists of a number of β strands that lie parallel to one another and are joined together by a regular array of hydrogen bonds between the carbonyl and imine groups of the neighboring backbones. Neighboring segments of the polypeptide backbone may lie either parallel (in the same N-terminal → C-terminal direction) or antiparallel (in the opposite N-terminal → C-terminal direction).
139. Tertiary Structure
140. Figure 2.32: A ribbon model of ribonuclease. The regions of α helix are depicted as spirals and β strands as flattened ribbons with the arrows indicating the N-terminal → C-terminal direction of the polypeptide. Those segments of the chain that do not adopt a regular secondary structure (i.e., an α helix or β strand) consist largely of loops and turns. Disulfide bonds are shown in yellow.
141. Figure 2.33: An X-ray diffraction pattern of myoglobin. The pattern of spots is produced as a beam of X-rays is diffracted by the atoms in the protein crystal, causing the X-rays to strike the film at specific sites. Information derived from the position and intensity (darkness) of the spots can be used to calculate the positions of the atoms in the protein that diffracted the beam, leading to complex structures such as that shown in Figure 2.34.
142. Myoglobin: The First Globular Protein Whose Tertiary Structure Was Determined
143. Figure 2.34: The three-dimensional structure of myoglobin. (a) The tertiary structure of whale myoglobin. Most of the amino acids are part of α helices. The nonhelical regions occur primarily as turns, where the polypeptide chain changes direction. The position of the heme is indicated in red. (b) The three-dimensional structure of myoglobin (heme indicated in red). The positions of all of the molecule’s atoms, other than hydrogen, are shown.
144. Protein Domains
145. Figure 2.35: Types of noncovalent bonds maintaining the conformation of proteins.
146. Figure 2.36: Proteins are built of structural units, or domains. The mammalian enzyme phospholipase C is constructed of four domains, indicated in different colors. The catalytic domain of the enzyme is shown in blue. Each of the domains of this enzyme can be found independently in other proteins as indicated by the matching color.
147. Dynamic Changes within Proteins
148. Figure 2.37: Dynamic movements within the enzyme acetylcholinesterase. A portion of the enzyme is depicted here in two different conformations: (1) a closed conformation (left) in which the entrance to the catalytic site is blocked by the presence of aromatic rings that are part of the side chains of tyrosine and phenylalanine residues (shown in purple) and (2) an open conformation (right) in which the aromatic rings of these side chains have swung out of the way, opening the “gate” to allow acetylcholine molecules to enter the catalytic site. These images are constructed using computer programs that take into account a host of information about the atoms that make up the molecule, including bond lengths, bond angles, electrostatic attraction and repulsion, van der Waals forces, etc. Using this information, researchers are able to simulate the movements of the various atoms over a very short time period, which provides images of the conformations that the protein can assume. An animation of this image can be found on the Web at http://mccammon.ucsd.edu (Courtesy of J. Andrew McCammon.)
149. Quaternary Structure
150. Protein-Protein Interactions
151. Figure 2.38: Proteins with quaternary structure. (a) Drawing of transforming growth factor-β2 (TGF-β2), a protein that is a dimer composed of two identical subunits. The two subunits are colored yellow and blue. Shown in white are the cysteine side chains and disulfide bonds. The spheres shown in yellow and blue are the hydrophobic residues that form the interface between the two subunits. (b) Drawing of a hemoglobin molecule, which consists of two α-globin chains and two β-globin chains (a heterotetramer) joined by noncovalent bonds. When the four globin polypeptides are assembled into a complete hemoglobin molecule, the kinetics of O2 binding and release are quite different from those exhibited by isolated polypeptides. This is because the binding of O2 to one polypeptide causes a conformational change in the other polypeptides that alters their affinity for O2 molecules.
152. Figure 2.39: Pyruvate dehydrogenase: a multiprotein complex. (a) Electron micrograph of a negatively stained pyruvate dehydrogenase complex isolated from E. coli. Each complex contains 60 polypeptide chains constituting three different enzymes. Its molecular mass approaches 5 million daltons. (b) A model of the pyruvate dehydrogenase complex. The core of the complex consists of a cube-like cluster of dihydrolipoyl transacetylase molecules. Pyruvate dehydrogenase dimers (black spheres) are distributed symmetrically along the edges of the cube, and dihydrolipoyl dehydrogenase dimers (small gray spheres) are positioned in the faces of the cube.
153. Figure 2.40: Protein–protein interactions. (a) A model illustrating the complementary molecular surfaces of portions of two interacting proteins. The reddish-colored molecule is an SH3 domain of the enzyme PI3K, whose function is discussed in Chapter 15. This domain binds specifically to a variety of proline-containing peptides, such as the one shown in the space-filling model at the top of the figure. The proline residues in the peptide, which fit into hydrophobic pockets on the surface of the enzyme, are indicated. The polypeptide backbone of the peptide is colored yellow, and the side chains are colored green. (b) Schematic model of the interaction between an SH3 domain and a peptide showing the manner in which certain residues of the peptide fit into hydrophobic pockets in the SH3 domain.
154. Figure 2.41: A network of protein-protein interactions. Each red line represents an interaction between two yeast proteins, which are indicated by the named black dots. In each case, the arrow points from an SH3 domain protein to a target protein with which it can bind. The 59 interactions depicted here were detected using two different types of techniques that measure protein-protein interactions. (See Trends Biochem. Sci. 34:1, and 579, 2009, for discussion of the validity of protein-protein interaction studies.)
155. Figure 2.42: Protein–protein interactions of hub proteins. (a) The enzyme RNA polymerase II, which synthesizes messenger RNAs in the cell, binds a multitude of other proteins simultaneously using multiple interfaces. (b) The enzyme Cdc28, which phosphorylates other proteins as it regulates the cell division cycle of budding yeast. Cdc28 binds a number of different proteins (Cln1-Cln3) at the same interface, which allows only one of these partners to bind at a time.
156. Protein Folding
157. Figure 2.43: Denaturation and refolding of ribonuclease. A native ribonuclease molecule (with intramolecular disulfide bonds indicated) is reduced and unfolded with β-mercaptoethanol and 8 M urea. After removal of these reagents, the protein undergoes spontaneous refolding. (From C. J., Epstein, R. F., Goldberger, and C. B., Anfinsen, Cold Spring Harbor Symp. Quant. Biol. 28:439, 1963. Reprinted with Permission from Cold Spring Harbor Laboratory Press.)
158. Figure 2.44: Two alternate pathways by which a newly synthesized or denatured protein could achieve its native conformation. Curled segments represent α helices, and arrows represent β strands.
159. Figure 2.45: Along the folding pathway. The image on the left shows the native tertiary structure of the enzyme acyl-phosphatase. The image on the right is the transition structure, which represents the state of the molecule at the top of an energy barrier that must be crossed if the protein is going to reach the native state. The transition structure consists of numerous individual lines because it is a set (ensemble) of closely related structures. The overall architecture of the transition structure is similar to that of the native protein, but many of the finer structural features of the fully folded protein have yet to emerge. Conversion of the transition state to the native protein involves completing secondary structure formation, tighter packing of the side chains, and finalizing the burial of hydrophobic side chains from the aqueous solvent.
160. The Role of Molecular Chaperones
161. Figure 2.46: The role of molecular chaperones in encouraging protein folding. The steps are described in the text. (Other families of chaperones are known but are not discussed.)
162. THE HUMAN PERSPECTIVE: Protein Misfolding Can Have Deadly Consequences
163. Figure 1: A contrast in structure. (a) Tertiary structure of the normal (PrPC) protein as determined by NMR spectroscopy. The orange portions represent α-helical segments, and the blue portions are short β strands. The yellow dotted line represents the N-terminal portion of the polypeptide, which lacks defined structure. (b) A proposed model of the abnormal, infectious (PrPSc) prion protein, which consists largely of β-sheet. The actual tertiary structure of the prion protein has not been determined. The two molecules shown in this figure are formed by polypeptide chains that can be identical in amino acid sequence but fold very differently. As a result of the differences in folding, PrPC remains soluble, whereas PrPSc produces aggregates that kill the cell. (The two molecules shown in this figure are called conformers because they differ only in conformation.)
164. Figure 2: Alzheimer’s disease. (a) The defining characteristics of brain tissue from a person who died of Alzheimer’s disease. (b) Amyloid plaques containing aggregates of the Aβ peptide appear extracellularly (between nerve cells), whereas neurofibrillary tangles (NFTs) appear within the cells themselves. NFTs, which are discussed at the end of the Human Perspective, are composed of misfolded tangles of a protein called tau that is involved in maintaining the microtubule organization of the nerve cell. Both the plaques and tangles have been implicated as a cause of the disease.
165. Figure 3: Formation of the A β peptide. The Aβ peptide is carved from the amyloid precursor protein (APP) as the result of cleavage by two enzymes, β-secretase and γ-secretase. It is interesting that APP and the two secretases are all proteins that span the membrane. Cleavage of APP occurs inside the cell (probably in the endoplasmic reticulum), and the Aβ product is ultimately secreted into the space outside of the cell. The γ-secretase can cut at either of two sites in the APP molecule, producing either Aβ 40 or Aβ42 peptides, the latter of which is primarily responsible for production of the amyloid plaques seen in Figure 2.48. γ-Secretase is a multisubunit enzyme that cleaves its substrate at a site within the membrane.
166. Figure 4: A neuroimaging technique that reveals the presence of amyloid in the brain. These PET (positron emission tomography) scans show the brains of two individuals that have ingested a radioactive compound, called Amyvid, that binds to amyloid deposits and appears red in the image. The top shows a healthy brain and the bottom a brain from a patient with AD, revealing extensive amyloid build-up. Amyloid deposits in the brain can be detected with this technique in persons who show no evidence of cognitive dysfunction. Such symptom-free individuals are presumed to be at high risk of going on to develop AD. Those who lack such deposits can be considered at very low risk of the disease in the near future.
167. The Emerging Field of Proteomics
168. Figure 2.47 : © Joseph G. Sutliff.
169. Figure 2.48: The study of proteomics often requires the separation of complex mixtures of proteins. The two electrophoretic gels shown here contain proteins extracted from the frontal cortex of humans (a) or chimpanzees (b). The numbered spots represent homologous proteins that show distinct differences in the two gels as discussed in the text.
170. Figure 2.49: Identifying proteins by mass spectrometry. A protein is isolated from a source (such as one of the spots on one of the gels of Figure 2.52) and subjected to digestion by the enzyme trypsin. The peptide fragments are then introduced into a mass spectrometer where they are ionized and separated according to their mass/charge (m/z) ratio. The separated peptides appear as a pattern of peaks whose precise m/z ratio is indicated. A comparison of these ratios to those obtained by a theoretical digest of virtual proteins encoded by the genome allows researchers to identify the protein being studied. In this case, the MS spectrum is that of horse myoglobin lacking its heme group.
171. FIGURE IN FOCUS
172. Figure 2.50: Global analysis of protein activities using protein chips. (a) This single microscope slide contains 5800 different yeast proteins spotted in duplicate. The proteins spotted on the slide were synthesized in genetically engineered cells. The spots display red fluorescence because they have been incubated with a fluorescent antibody that can bind to all of the proteins in the array. (b) The left image shows a small portion of the protein array depicted in part a. The right image shows the same portion of the array following incubation with calmodulin in the presence of calcium ions. The two proteins exhibiting green fluorescence are calmodulin-binding proteins. (c) Schematic illustration of the events occurring in part b where calmodulin has bound to two proteins of the microarray that have complementary binding sites.
173. Protein Engineering
174. Figure 2.51: The computational design of a protein that is capable of binding specifically to the surface of another protein. (a) The computationally designed protein is shown in green and its target protein (the HA protein from the H1N1 1918 influenza virus) is shown on the left in multiple colors. The predicted structure of the designed protein fits closely with that of the actual protein that was generated from the predicted sequence. (b) The actual interface of the targeted hydrophobic helix of the HA protein (gray) and the designed protein (purple). Side chains of the designed protein are seen to interact with sites on the HA helix.
175. Structure-Based Drug Design
176. Figure 2.52: Development of a protein-targeting drug, such as Gleevec. (a) Typical steps in drug development. In step 1, a protein (e.g., ABL) has been identified that plays a causative role in the disease. This protein is a likely target for a drug that inhibits its enzymatic activity. In step 2, the protein is incubated with thousands of compounds in a search for ones that bind with reasonable affinity and inhibit its activity. In step 3, one such compound (e.g, 2-phenylaminopyrimidine in the case of ABL) has been identified. In step 4, knowledge of the structure of the target protein is used to make derivatives of the compound (e.g., Gleevec) that have greater binding affinity and thus can be used at lower concentrations. In step 5, the compound in question is tested in preclinical experiments for toxicity and efficacy (level of effectiveness) in vivo. Preclinical experiments are typically carried out on cultured human cells (step 5a) (e.g., those from patients with CML) and laboratory animals (step 5b) (e.g., mice carrying transplants of human CML cells). If the drug appears safe and effective in animals, the drug is tested in clinical trials (step 6) as discussed on page 68. (b) The structure of Gleevec. The blue portion of the molecule indicates the structure of the compound 2-phenylaminopyrimidine that was initially identified as an ABL kinase inhibitor. (c,d) The structure of the catalytic domain of ABL in complex (c) with Gleevec (shown in yellow) and (d) with a second-generation inhibitor called Sprycel. Gleevec binds to the inactive conformation of the protein, whereas Sprycel binds to the active conformation. Both binding events block the activity that is required for the cell’s cancerous phenotype. Sprycel is effective against most cancer cells that have become resistant to the action of Gleevec.
177. Protein Adaptation and Evolution
178. Figure 2.53: Distribution of polar, charged amino acid residues in the enzyme malate dehydrogenase from a halophilic archaebacterium. Red balls represent acidic residues, and blue balls represent basic residues. The surface of the enzyme is seen to be covered with acidic residues, which gives the protein a net charge of −156, and promotes its solubility in extremely salty environments. For comparison, a homologous protein from the dogfish, an ocean-dwelling shark, has a net charge of +16.
179. Figure 2.54: The dramatic effect on conformation that can result from a single amino acid substitution. In this case the switch between a leucine and a tyrosine at a critical position within this 56-amino acid polypeptide chain results in a transformation of the entire fold of the backbone of this polypeptide. This single substitution causes 85 percent of the amino acid residues to change their secondary structure. The spatial disposition of the two alternate side chains, which brings about this conformational shift, is shown in red in the model structures. The N-terminal amino acids are shown in orange and the C-terminal amino acids in blue.
180. Nucleic Acids
181. Figure 2.55: Nucleotides and nucleotide strands of RNA.(a) Nucleotides are the monomers from which strands of nucleic acid are constructed. A nucleotide consists of three parts: a sugar, a nitrogenous base, and a phosphate. The nucleotides of RNA contain the sugar ribose, which has a hydroxyl group bonded to the second carbon atom. In contrast, the nucleotides of DNA contain the sugar deoxyribose, which has a hydrogen atom rather than a hydroxyl group attached to the second carbon atom. Each nucleotide is polarized, having a 5′ end (corresponding to the 5′ side of the sugar) and a 3′ end. (b) Nucleotides are joined together to form a strand by covalent bonds that link the 3′ hydroxyl group of one sugar with the 5′ phosphate group of the adjoining sugar.
182. Figure 2.56: Nitrogenous bases in nucleic acids. Of the four standard bases found in RNA, adenine and guanine are purines, and uracil and cytosine are pyrimidines. In DNA, the pyrimidines are cytosine and thymine, which differs from uracil by a methyl group attached to the ring.
183. Figure 2.57: RNAs can assume complex shapes. (a) This ribosomal RNA is an integral component of the small ribosomal subunit of a bacterium. In this two-dimensional profile, the RNA strand is seen to be folded back on itself in a highly ordered pattern so that most of the molecule is double-stranded. (b) This hammerhead ribozyme, as it is called, is a small RNA molecule from a viroid (page 26). The helical nature of the double-stranded portions of this RNA can be appreciated in this three-dimensional model of the molecule.
184. REVIEW
185. 2.6: The Formation of Complex Macromolecular Structures
186. The Assembly of Tobacco Mosaic Virus Particles and Ribosomal Subunits
187. Figure 2.58: Reconstruction of a ribosome from the cytoplasm of a wheat germ cell. This reconstruction is based on high-resolution electron micrographs and shows the two subunits of this eukaryotic ribosome, the small (40S) subunit on the left and the large (60S) subunit on the right. The internal structure of a ribosome is discussed in Section 11.8.
188. REVIEW
189. EXPERIMENTAL PATHWAYS: Chaperones: Helping Proteins Reach Their Proper Folded State
190. Figure 1: A model of the GroEL complex built according to data from electron microscopy and molecular-weight determination. The complex is seen to consist of two disks, each composed of seven identical subunits arranged symmetrically around a central axis. Subsequent studies showed the complex contains two internal chambers.
191. Figure 2: Reconstructions of GroEL based on high-resolution electron micrographs taken of specimens that had been frozen in liquid ethane and examined at −170° C. The image on the left shows the GroEL complex, and that on the right shows the GroEL complex with GroES, which appears as a dome on one end of the cylinder. It is evident that the binding of the GroES is accompanied by a marked change in conformation of the apical end of the proteins that make up the top GroEL ring (arrow), which results in a marked enlargement of the upper chamber.
192. Figure 3 Conformational change in GroEL. (a) The model on the left shows a surface view of the two rings that make up the GroEL chaperonin. The drawing on the right shows the tertiary structure of one of the subunits of the top GroEL ring. The polypeptide chain can be seen to fold into three domains. (b) When a GroES ring (arrow) binds to the GroEL cylinder, the apical domain of each GroEL subunit of the adjacent ring undergoes a dramatic rotation of approximately 60° with the intermediate domain (shown in green) acting like a hinge. The effect of this shift in parts of the polypeptide is a marked elevation of the GroEL wall and enlargement of the enclosed chamber.
193. Figure 4: A schematic illustration of the proposed steps that occur during the GroEL-GroES-assisted folding of a polypeptide. The GroEL is seen to consist of two chambers that have equivalent structures and functions and that alternate in activity. Each chamber is located within one of the two rings that make up the GroEL complex. The nonnative polypeptide enters one of the chambers (step 1) and binds to hydrophobic sites on the chamber wall. Binding of the GroES cap produces a conformational change in the wall of the top chamber, causing the enlargement of the chamber and release of the nonnative polypeptide from the wall into the encapsulated space (step 2). After about 15 seconds have elapsed, the GroES dissociates from the complex and the polypeptide is ejected from the chamber (step 3). If the polypeptide has achieved its native conformation, as has the molecule on the left, the folding process is complete. If, however, the polypeptide is only partially folded, or is misfolded, it will rebind the GroEL chamber for another round of folding.
194. References
195. Synopsis
196. Analytic Questions
197. 3: Bioenergetics, Enzymes, and Metabolism
198. A model showing the surface of the enzyme Δ5-3-ketosteroid isomerase with a substrate molecule (green) in the active site. The electrostatic character of the surface is indicated by color (red, acidic; blue, basic).
199. 3.1: Bioenergetics
200. The Laws of Thermodynamics and the Concept of Entropy
201. The First Law of Thermodynamics
202. Figure 3.1: Examples of energy transduction. (a) Conversion of electrical energy to mechanical energy, (b) conversion of chemical energy to mechanical and thermal energy, (c) conversion of chemical energy to light energy.
203. Figure 3.2: A change in a system’s internal energy. In this example, the system will be defined as a particular leaf of a plant. (a) During the day, sunlight is absorbed by photosynthetic pigments in the leaf’s chloroplasts and used to convert CO2 into carbohydrates, such as the glucose molecule shown in the drawing (which is subsequently incorporated into sucrose or starch). As the cell absorbs light, its internal energy increases; the energy present in the remainder of the universe has to decrease. (b) At night, the energy relationship between the cell and its surroundings is reversed as the carbohydrates produced during the day are oxidized to CO2 in the mitochondria and the energy is used to run the cell’s nocturnal activities.
204. The Second Law of Thermodynamics
205. Figure 3.3: Events are accompanied by an increase in the entropy of the universe. (a) A sugar cube contains sucrose molecules in a highly ordered arrangement in which their freedom of movement is restricted. As the cube dissolves, the freedom of movement of the sucrose molecules is greatly increased, and their random movement causes them to become equally distributed throughout the available space. Once this occurs, there will be no further tendency for redistribution, and the entropy of the system is at a maximum. (b) Sugar molecules spread randomly through a solution can be returned to an ordered state, but only if the entropy of the surroundings is increased, as occurs when the more ordered water molecules of the liquid phase become disordered following evaporation.
206. Free Energy
207. Figure 3.4: When water freezes, its entropy decreases because the water molecules of ice exist in a more ordered state with less freedom of movement than in a liquid state. The decrease in entropy is particularly apparent in the formation of a snowflake.
208. Free-Energy Changes in Chemical Reactions
209. Table 3.1: Thermodynamics of the Ice–Water Transformation
210. Table 3.2: Relation between ΔG°′ and Keq′ at 25°C
211. Free-Energy Changes in Metabolic Reactions
212. Figure 3.5: ATP hydrolysis. Adenosine triphosphate (ATP) is hydrolyzed as part of many biochemical processes. In most reactions, as shown here, ATP is hydrolyzed to ADP and inorganic phosphate (Pi), but in some cases (not shown) it is hydrolyzed to AMP, a compound with only one phosphate group, and pyrophosphate (PPi). Both of these reactions have essentially the same ΔG°′ of −7.3 kcal/mol (−30.5 kJ/mol).
213. Coupling Endergonic and Exergonic Reactions
214. FIGURE IN FOCUS
215. Figure 3.6: A few roles for ATP hydrolysis. In the cell, energy from ATP hydrolysis may be used to (a) separate charge across a membrane; (b) concentrate a particular solute within the cell; (c) drive an otherwise unfavorable chemical reaction; (d) slide filaments across one another, as occurs during the shortening of a muscle cell; (e) donate a phosphate group to a protein, thereby changing its properties and bringing about a desired response. In this case the added phosphate groups serve as binding sites for other proteins.
216. Equilibrium versus Steady-State Metabolism
217. Figure 3.7: Steady state versus equilibrium. (a) As long as this ameba can continue to take in nutrients from the outside world, it can harvest the energy necessary to maintain concentrations of compounds at a steady state, which may be far from equilibrium. The steady-state concentrations of ATP and ADP are indicated by the colored dots and the histogram. (b) When the ameba dies, the concentrations of ATP and ADP (as well as other biochemicals) shift toward their equilibrium ratios.
218. REVIEW
219. 3.2: Enzymes as Biological Catalysts
220. The Properties of Enzymes
221. Table 3.3: Catalytic Activity of a Variety of Enzymes
222. Overcoming the Activation Energy Barrier
223. Figure 3.8: Activation energy and enzymatic reactions. Even though the formation of glucose 6-phosphate is a thermodynamically favored reaction (ΔG°′=−4 kcal/mol), the reactants must possess sufficient energy to achieve an activated state in which the atomic rearrangements necessary for the reaction can occur. The amount of energy required is called the activation energy (EA) and is represented by the height of the curve. The activation energy is not a fixed value, but varies with the particular reaction pathway. EA is greatly reduced when the reactants combine with an enzyme catalyst. (This diagram depicts a simple, one-step reaction mechanism. Many enzymatic reactions take place in two or more steps leading to the formation of intermediates (as in Figure 3.13). Each step in the reaction has a distinct EA and a separate transition state.)
224. Figure 3.9: The effect of lowering activation energy on the rate of a reaction. The bell-shaped curves indicate the energy content of a population of molecules present in a reaction mixture at two different temperatures. The number of reactant molecules containing sufficient energy to undergo reaction is increased by either heating the mixture or adding an enzyme catalyst. Heat increases the rate of reaction by increasing the energy content of the molecules, while the enzyme does so by lowering the activation energy required for the reaction to occur.
225. Figure 3.10: Formation of an enzyme–substrate complex. Schematic drawing of the reaction catalyzed by pyruvate kinase (see Figure 3.24) in which the two substrates, phosphoenolpyruvate (PEP) and ADP, bind to the enzyme to form an enzyme–substrate (ES) complex, which leads to the formation of the products, pyruvate and ATP.
226. The Active Site
227. Figure 3.11: The active site of an enzyme. (a) Diagrammatic representation of the active site of the enzyme ribulose bisphosphate carboxylase oxygenase showing the various sites of interaction between the bound substrates (RuBP and CO2) and certain amino acid side chains of the enzyme. In addition to determining the substrate-binding properties of the active site, these noncovalent interactions alter the properties of the substrate in ways that accelerate its conversion to products. (b) An electron density map of the active site of a viral thymidine kinase with the substrate, deoxythymidine, shown in the center of the map (arrow). The blue mesh provides an indication of the outer reaches of the electron orbitals of the atoms that make up the substrate and the side chains of the enzyme, thus portraying a visual representation of the space occupied by the atoms of the active site. (c,d) Examples of the precise fit that occurs between parts of an enzyme and a substrate during catalysis. These two examples show the tight spatial relationship between (c) a glutamic acid (yellow) and (d) a histidine (green) of the enzyme triosephosphate isomerase and the substrate (red).
228. Mechanisms of Enzyme Catalysis
229. Substrate Orientation
230. Changing Substrate Reactivity
231. Figure 3.12: Three mechanisms by which enzymes accelerate reactions: (a) maintaining precise substrate orientation, (b) changing substrate reactivity by altering its electrostatic configuration, (c) exerting physical stress on bonds in the substrate to be broken.
232. Figure 3.13: Diagrammatic representation of the catalytic mechanism of chymotrypsin. The reaction is divided into two steps. (a) The electronegative oxygen atom of a serine residue (Ser 195) in the enzyme, which carries a partial negative charge, carries out a nucleophilic attack on the carbonyl carbon atom of the substrate, which carries a partial positive charge, splitting the peptide bond. The polypeptide substrate is shown in blue. The serine is made more reactive by a closely applied histidine residue (His 57) that draws the proton from the serine and subsequently donates the proton to the nitrogen atom of the cleaved peptide bond. Histidine is able to do this because its side chain is a weak base that is capable of gaining and losing a proton at physiologic pH. (A stronger base, such as lysine, would remain fully protonated at this pH.) Part of the substrate forms a transient covalent bond with the enzyme by means of the serine side chain, while the remainder of the substrate is released as the first product. (It can be noted that the serine and histidine residues are situated 138 amino acids away from each other in the primary sequence but are brought together within the enzyme by the folding of the polypeptide. An aspartic acid, residue 102, which is not shown, also plays a role in catalysis by influencing the ionic state of the histidine.) (b) In the second step, the electronegative oxygen atom of a water molecule displaces the covalently linked substrate from the enzyme, regenerating the unbound enzyme molecule. As in the first step, the histidine plays a role in proton transfer; in this step, the proton is removed from water, making it a much stronger nucleophile. The proton is subsequently donated to the serine residue of the enzyme.
233. Inducing Strain in the Substrate
234. Figure 3.14: An example of induced fit. When a glucose molecule binds to the enzyme hexokinase, the protein undergoes a conformational change that encloses the substrate within the active site pocket and aligns the reactive groups of the enzyme and substrate.
235. Figure 3.15: Electron density map of a nsingle hydrogen bond (green-dotted line). This map shows a very small part of the proteolytic enzyme subtilisin at atomic (0.78 Å) resolution. The hydrogen atom (yellow) is seen to be shared between a nitrogen atom on the ring of a histidine residue and an oxygen atom of an aspartic acid residue.
236. Figure 3.16: Myoglobin: the movie. In this example of time-resolved X-ray crystallography, the structure of myoglobin (Mb) was determined with a molecule of CO bound to the heme group and at various times after the CO molecule was released. (CO binds to the same site on Mb as O2 but is better suited for these types of studies.) Release of CO was induced simultaneously throughout the crystal by exposure to a flash of laser light (photolysis). Each of the structures was determined following a single, intense X-ray pulse from a synchrotron. The myoglobin molecule being studied had a single amino acid substitution that made it a better subject for analysis. (a) A 6.5 Å-thick slice through a Mb molecule showing the changes that occur by 100 picoseconds (1 ps=one-trillionth of a second) following the release of CO from its binding site. The structure of Mb prior to the laser flash is shown in magenta, and the structure of the protein 100 ps after the laser flash is shown in green. Those parts of the molecule that did not change in structure over this time period appear white. Three large-scale displacements near the CO-binding site can be seen (indicated by the yellow arrows). (b) An enlarged view of the boxed region in part a. The released CO (solid circle) is situated about 2 Å from its original binding site (dashed circle). Movement of CO is accommodated by the rotation of Phe29, which pushes His64 outward, which in turn dislodges a bound water molecule. (c) By 3.16 nanoseconds after the laser flash, the CO molecule has migrated to either of the two positions shown (labeled 2 and 3), Phe29 and His64 have relaxed toward their initial states, and the heme group has undergone considerable displacement as indicated by the increased amount of green shading in the region of the heme.
237. Enzyme Kinetics
238. Figure 3.17: The relationship between the rate (velocity) of an enzyme-catalyzed reaction and the substrate concentration. Since each enzyme molecule is only able to catalyze a certain number of reactions in a given amount of time, the velocity of the reaction (typically expressed as moles of product formed per second) approaches a maximal rate as the substrate concentration increases. The substrate concentration at which the reaction is at half-maximal velocity (Vmax/2) is called the Michaelis constant, or KM.
239. Figure 3.18: A Lineweaver-Burk plot of the reciprocals of velocity and substrate concentration from which the values for the Vmax and KM are readily calculated.
240. Figure 3.19: Dependence of the rate of an enzyme-catalyzed reaction on (a) pH, and (b) temperature. The shape of the curves and the optimal pH and temperature vary with the particular reaction. (a) Changes in pH affect the ionic properties of the substrate and the enzyme as well as the enzyme’s conformation. (b) At lower temperatures, reaction rates rise with increasing temperature due to the increased energy of the reactants. At higher temperatures, this positive effect is offset by enzyme denaturation.
241. Enzyme Inhibitors
242. Figure 3.20: Competitive inhibition. Because of their molecular similarity, competitive inhibitors are able to compete with the substrate for a binding site on the enzyme. The effect of a competitive inhibitor depends on the relative concentrations of the inhibitor and substrate.
243. REVIEW
244. Figure 3.21: The effects of inhibitors on enzyme kinetics. The effect of both competitive and noncompetitive inhibitors is shown when the kinetics of the reaction are plotted as velocity of reaction versus substrate concentration (a) or its reciprocal (b). The noncompetitive inhibitor reduces Vmax without affecting KM, whereas the competitive inhibitor increases KM without affecting Vmax. (The more complex cases of uncompetitive and mixed inhibitors are not discussed.)
245. THE HUMAN PERSPECTIVE: The Growing Problem of Antibiotic Resistance
246. Table 1: Antibiotics in Clinical Use and Modes of Resistance
247. 3.3: Metabolism
248. An Overview of Metabolism
249. Figure 3.22: Three stages of metabolism. The catabolic pathways (green arrows rightward) converge to form common metabolites and lead to ATP synthesis in stage III. The anabolic pathways (blue arrows leftward) start from a few precursors in stage III and utilize ATP to synthesize a large variety of cellular materials. Metabolic pathways for nucleic acids are more complex and are not shown here.
250. Oxidation and Reduction: A Matter of Electrons
251. Figure 3.23: The oxidation state of a carbon atom depends on the other atoms to which it is bonded. Each carbon atom can form a maximum of four bonds with other atoms. This series of simple, one-carbon molecules illustrates the various oxidation states in which the carbon atom can exist. In its most reduced state, the carbon is bonded to four hydrogens (forming methane); in its most oxidized state, the carbon atom is bonded to two oxygens (forming carbon dioxide).
252. The Capture and Utilization of Energy
253. Glycolysis and ATP Formation
254. Figure 3.24: The steps of glycolysis.
255. Figure 3.25: Free-energy profile of glycolysis in cardiac muscle tissue. The step numbers correspond to the reactions of glycolysis shown in Figure 3.24. All reactions are at or near equilibrium except those catalyzed by hexokinase, phosphofructokinase, and pyruvate kinase (reactions 1, 3, and 10), which exhibit large differences in free energy.
256. Figure 3.26: The transfer of energy during a chemical oxidation. The oxidation of glyceraldehyde 3-phosphate to 3-phosphoglycerate, which is an example of the oxidation of an aldehyde to a carboxylic acid, occurs in two steps catalyzed by two enzymes. The first reaction (parts a and b) is catalyzed by the enzyme glyceraldehyde 3-phosphate dehydrogenase, which transfers a hydride ion (two electrons and a proton) from the substrate to NAD+. Once reduced, the NADH molecules are displaced by NAD+ molecules from the cytosol (a) and the bound substrate is phosphorylated and released (b). The second reaction (part c), which is catalyzed by the enzyme phosphoglycerate kinase, is an example of a substrate-level phosphorylation in which a phosphate group is transferred from a substrate molecule, in this case 1,3-bisphosphoglycerate, to ADP to form ATP.
257. Figure 3.27: The structure of NAD+ and its reduction to NADH. When the 2′ OH of the ribose moiety (indicated by the purple colored screen) is covalently bonded to a phosphate group, the molecule is NADP+/NADPH, whose function is discussed later in the chapter.
258. Figure 3.28: Ranking compounds by phosphate transfer potential. Those phosphorylated compounds higher on the scale (ones with a more negative ΔG°′ of hydrolysis) have a lower affinity for their phosphate group than those compounds lower on the scale. As a result, compounds higher on the scale readily transfer their phosphate group to form compounds that are lower on the scale. Thus, phosphate groups can be transferred from 1,3-bisphosphate or phosphoenolpyruvate to ADP during glycolysis.
259. Anaerobic Oxidation of Pyruvate: The Process of Fermentation
260. Figure 3.29: Fermentation. Most cells carry out aerobic respiration, which depends on molecular oxygen. If oxygen supplies should diminish, as occurs in a skeletal muscle cell undergoing strenuous contraction or a yeast cell living under anaerobic conditions, these cells are able to regenerate NAD+ by fermentation. Muscle cells accomplish fermentation by formation of lactate, whereas yeast cells do so by formation of ethanol. Aerobic oxidation of pyruvate by means of the TCA cycle is discussed at length in Chapter 5.
261. Reducing Power
262. Metabolic Regulation
263. Altering Enzyme Activity by Covalent Modification
264. Altering Enzyme Activity by Allosteric Modulation
265. Separating Catabolic and Anabolic Pathways
266. Figure 3.30: Feedback inhibition. The flow of metabolites through a metabolic pathway stops when the first enzyme of the pathway (enzyme BC) is inhibited by the end product of that pathway (compound E), which binds to an allosteric site on the enzyme. Feedback inhibition prevents a cell from wasting resources by continuing to produce compounds that are no longer required.
267. Figure 3.31: Glycolysis versus gluconeogenesis. Whereas most of the reactions are the same in the two pathways, even though they run in opposite directions, the three irreversible reactions of glycolysis (steps 1 to 3 here) are replaced in the gluconeogenic pathway by different, thermodynamically favored reactions.
268. REVIEW
269. Synopsis
270. Analytic Questions
271. 4: The Structure and Function of the Plasma Membrane
272. Three-dimensional, X-ray crystallographic structure of a β2-adrenergic receptor (β2-AR), which is a member of the G protein-coupled receptor (GPCR) superfamily. These integral membrane proteins are characterized as containing seven transmembrane helices. The β2-AR is a resident of the plasma membrane of a variety of cells, where it normally binds the ligand epinephrine and mediates such responses as increased heart rate and relaxation of smooth muscle cells. Until recently, GPCRs had been very difficult to crystallize so that high-resolution structures of these important proteins have been lacking. This situation is now rapidly changing as the result of recent advances in crystallization technology. The image shown here depicts two β2-ARs, which were crystallized in the presence of cholesterol and palmitic acid (yellow) and a receptor-binding ligand (green). The crystals used to obtain this image were selected from more than 15,000 trials.
273. Figure 4.1: The trilaminar appearance of membranes. (a) Electron micrograph showing the three-layered (trilaminar) structure of the plasma membrane of an erythrocyte after staining the tissue with the heavy metal osmium. Osmium binds preferentially to the polar head groups of the lipid bilayer, producing the trilaminar pattern. The arrows denote the inner and outer edges of the membrane. (b) The outer edge of a differentiated muscle cell grown in culture showing the similar trilaminar structure of both the plasma membrane (PM) and the membrane of the sarcoplasmic reticulum (SR), a calcium-storing compartment of the cytoplasm.
274. 4.1: An Overview of Membrane Functions
275. Figure 4.2: A summary of membrane functions in a plant cell. (1) An example of membrane compartmentalization in which hydrolytic enzymes (acid hydrolases) are sequestered within the membrane-bounded vacuole. (2) An example of the role of cytoplasmic membranes as a site of enzyme localization. The fixation of CO2 by the plant cell is catalyzed by an enzyme that is associated with the outer surface of the thylakoid membranes of the chloroplasts. (3) An example of the role of membranes as a selectively permeable barrier. Water molecules are able to penetrate rapidly through the plasma membrane, causing the plant cell to fill out the available space and exert pressure against its cell wall. (4) An example of solute transport. Hydrogen ions, which are produced by various metabolic processes in the cytoplasm, are pumped out of plant cells into the extracellular space by a transport protein located in the plasma membrane. (5) An example of the involvement of a membrane in the transfer of information from one side to another (signal transduction). In this case, a hormone (e.g., abscisic acid) binds to the outer surface of the plasma membrane and triggers the release of a chemical message (such as IP3) into the cytoplasm. In this case, IP3 causes release of Ca2+ ions from a cytoplasmic warehouse. (6) An example of the role of membranes in cell–cell communication. Openings between adjoining plant cells, called plasmodesmata, allow materials to move directly from the cytoplasm of one cell into its neighbors. (7) An example of the role of membranes in energy transduction. The conversion of ADP to ATP occurs in close association with the inner membrane of the mitochondrion.
276. 4.2: A Brief History of Studies on Plasma Membrane Structure
277. Figure 4.3: The plasma membrane contains a lipid bilayer. (a) Calculating the surface area of a lipid preparation. When a sample of phospholipids is dissolved in an organic solvent, such as hexane, and spread over an aqueous surface, the phospholipid molecules form a layer over the water that is a single molecule thick: a monomolecular layer. The molecules in the layer are oriented with their hydrophilic groups bonded to the surface of the water and their hydrophobic chains directed into the air. To estimate the surface area the lipids would cover if they were part of a membrane, the lipid molecules can be compressed into the smallest possible area by means of movable barriers. Using this type of apparatus, which is called a Langmuir trough after its inventor, Gorter and Grendel concluded that red blood cells contained enough lipid to form a layer over their surface that was two molecules thick: a bilayer. (b) As Gorter and Grendel first proposed, the core of a membrane contains a bimolecular layer of phospholipids oriented with their water-soluble head groups facing the outer surfaces and their hydrophobic fatty acid tails facing the interior. The structures of the head groups are given in Figure 4.6a. (c) Simulation of a fully hydrated lipid bilayer composed of the phospholipid phosphatidylcholine. Phospholipid head groups are orange, water molecules are blue and white, fatty acid chains are green.
278. Figure 4.4: A brief history of the structure of the plasma membrane. (a) A revised 1954 version of the Davson-Danielli model showing the lipid bilayer, which is lined on both surfaces by a monomolecular layer of proteins that extends through the membrane to form protein-lined pores. (b) The fluid-mosaic model of membrane structure as initially proposed by Singer and Nicolson in 1972. Unlike previous models, the proteins penetrate the lipid bilayer. Although the original model shown here depicted a protein that was only partially embedded in the bilayer, lipid-penetrating proteins that have been studied span the entire bilayer. (c) A current representation of the plasma membrane showing the same basic organization as that proposed by Singer and Nicolson. The external surface of most membrane proteins, as well as a small percentage of the phospholipids, contain short chains of sugars, making them glycoproteins and glycolipids. Those portions of the polypeptide chains that extend through the lipid bilayer typically occur as α helices composed of hydrophobic amino acids. The two leaflets of the bilayer contain different types of lipids as indicated by the differently colored head groups. (d) Molecular model of the membrane of a synaptic vesicle constructed using known structures of the various proteins along with information on their relative numbers obtained from the analysis of purified synaptic vesicles. The high protein density of the membrane is apparent. Most of the proteins in this membrane are required for the interaction of the vesicle with the plasma membrane. The large blue protein at the lower right pumps H+ ions into the vesicle.
279. REVIEW
280. 4.3: The Chemical Composition of Membranes
281. Figure 4.5: The myelin sheath. Electron micrograph of a nerve cell axon surrounded by a myelin sheath consisting of concentric layers of plasma membrane that have an extremely low protein/lipid ratio. The myelin sheath insulates the nerve cell from the surrounding environment, which increases the velocity at which impulses can travel along the axon (discussed on page 167). The perfect spacing between the layers is maintained by interlocking protein molecules (called P0) that project from each membrane.
282. Membrane Lipids
283. Phosphoglycerides
284. Figure 4.6: The chemical structures of membrane lipids. (a) The structures of phosphoglycerides (see also Figure 2.22). (b) The structures of sphingolipids. Sphingomyelin is a phospholipid; cerebrosides and gangliosides are glycolipids. A third membrane lipid is cholesterol, which is shown in the next figure. (R=fatty acyl chain). [The green portion of each lipid, which represents the hydrophobic tail(s) of the molecule, is actually much longer than the hydrophilic head group (see Figure 4.23).]
285. Sphingolipids
286. Cholesterol
287. The Nature and Importance of the Lipid Bilayer
288. Figure 4.7: The cholesterol molecules (shown in green) of a lipid bilayer are oriented with their small hydrophilic end facing the external surface of the bilayer and the bulk of their structure packed in among the fatty acid tails of the phospholipids. The placement of cholesterol molecules interferes with the flexibility of the lipid hydrocarbon chains, which tends to stiffen the bilayer while maintaining its overall fluidity. Unlike other lipids of the membrane, cholesterol is often rather evenly distributed between the two layers (leaflets).
289. Table 4.1: Lipid Compositions of Some Biological Membranes*
290. Figure 4.8: The dynamic properties of the plasma membrane. (a) The leading edge of a moving cell often contains sites where the plasma membrane displays undulating ruffles. (b) Division of a cell is accompanied by the deformation of the plasma membrane as it is pulled toward the center of the cell. Unlike most dividing cells, the cleavage furrow of this dividing ctenophore egg begins at one pole and moves unidirectionally through the egg. (c) Membranes are capable of fusing with other membranes. This electron micrograph shows a secretory granule discharging its contents after fusion with the overlying plasma membrane (arrows).
291. The Asymmetry of Membrane Lipids
292. Figure 4.9: Liposomes. A schematic diagram of a stealth liposome containing a hydrophilic polymer (such as polyethylene glycol) to protect it from destruction by immune cells, antibody molecules that target it to specific body tissues, a water-soluble drug enclosed in the fluid-filled interior chamber, and a lipid-soluble drug in the bilayer.
293. Membrane Carbohydrates
294. Figure 4.10: The asymmetric distribution of phospholipids (and cholesterol) in the plasma membrane of human erythrocytes.
295. Figure 4.11: Two types of linkages that join sugars to a polypeptide chain. The N-glycosidic linkage between asparagine and N-acetylglucosamine is more common than the O-glycosidic linkage between serine or threonine and N-acetylgalactosamine.
296. Figure 4.12: Blood-group antigens. Whether a person has type A, B, AB, or O blood is determined by a short chain of sugars covalently attached to membrane lipids and proteins of the red blood cell membrane. The oligosaccharides attached to membrane lipids (forming a ganglioside) that produce the A, B, and O blood types are shown here. A person with type AB blood has gangliosides with both the A and B structure.
297. REVIEW
298. 4.4: The Structure and Functions of Membrane Proteins
299. Integral Membrane Proteins
300. Figure 4.13: Three classes of membrane protein. (a) Integral proteins typically contain one or more transmembrane helices (see Figure 5.4 for an exception). (b) Peripheral proteins are noncovalently bonded to the polar head groups of the lipid bilayer and/or to an integral membrane protein. (c) Lipid-anchored proteins are covalently bonded to a lipid group that resides within the membrane. The lipid can be phosphatidylinositol, a fatty acid, or a prenyl group (a long-chain hydrocarbon built from five-carbon isoprenoid units).
301. Figure 4.14: The interactions between membrane proteins and lipids. (a) Aquaporin is a membrane protein containing four subunits (colored differently in the illustration) with each subunit containing an aqueous channel. Analysis of the protein’s structure revealed the presence of a surrounding layer of bound lipid molecules. In this case, these lipid molecules are not likely to play a role in the function of aquaporin because the protein retains its function as a water channel in bilayers containing nonnative lipids. (b) Two views of another tetrameric membrane protein, in this case the bacterial K+ channel, KcsA. Anionic phosphatidylglycerol molecules (red/gray) are seen in each crevice between the subunits and are thought to be required for normal channel function. A K+ ion (purple sphere) is seen in transit through the pore.
302. Distribution of Integral Proteins: Freeze-Fracture Analysis
303. Figure 4.15: Freeze fracture: a technique for investigating cell membrane structure. (a) When a block of frozen tissue is struck by a knife blade, a fracture plane runs through the tissue, often following a path that leads it through the middle of the lipid bilayer. The fracture plane goes around the proteins rather than cracking them in half, and they segregate with one of the two halves of the bilayer. The exposed faces within the center of the bilayer can then be covered with a metal deposit to form a metallic replica. These exposed faces are referred to as the E, or ectoplasmic face, and the P, or protoplasmic face. (b) Replica of a freeze-fractured human erythrocyte. The P fracture face is seen to be covered with particles approximately 8 nm in diameter. A small ridge (arrow) marks the junction of the particulate face with the surrounding ice. (c) This micrograph shows the surface of an erythrocyte that was frozen and then fractured, but rather than preparing a replica, the cell was thawed, fixed, and labeled with a marker for the carbohydrate groups that project from the external surface of the integral protein glycophorin (Figure 4.18). Thin sections of the labeled, fractured cell reveal that glycophorin molecules (black particles) have preferentially segregated with the outer half of the membrane. The red line shows the path of the fracture plane.
304. Studying the Structure and Properties of Integral Membrane Proteins
305. Figure 4.16: Solubilization of membrane proteins with detergents. The nonpolar ends of the detergent molecules associate with the nonpolar residues of the protein that had previously been in contact with the fatty acyl chains of the lipid bilayer. In contrast, the polar ends of the detergent molecules interact with the surrounding water molecules, keeping the protein in solution. Nonionic detergents, as shown here, solubilize membrane proteins without disrupting their structure.
306. Figure 4.17: An integral protein as it resides within the plasma membrane. Tertiary structure of the photosynthetic reaction center of a bacterium as determined by X-ray crystallography. The protein contains three different membrane-spanning polypeptides, shown in yellow, light blue, and dark blue. The helical nature of each of the transmembrane segments is evident.
307. Identifying Transmembrane Domains
308. Figure 4.18: Glycophorin A, an integral protein with a single transmembrane domain. The single α helix that passes through the membrane consists predominantly of hydrophobic residues (brown-colored circles). The four positively charged amino acid residues of the cytoplasmic domain of the membrane protein form ionic bonds with negatively charged lipid head groups. Carbohydrates are attached to a number of amino acid residues on the outer surface of the protein (shown in the inset). All but one of the 16 oligosaccharides are small O-linked chains (the exception is a larger oligosaccharide linked to the asparagine residue at position 26). Glycophorin molecules are present as homodimers within the erythrocyte membrane (Figure 4.32d). The two helices of the dimer cross over one another in the region between residues 79 and 83. This Gly-X-X-X-Gly sequence is commonly found where transmembrane helices come into close proximity.
309. Figure 4.19: Accommodating various amino acid residues within transmembrane helices. (a) In this portrait of a small portion of a transmembrane helix, the hydroxyl group of the threonine side chain (arrow) is able to form a (shared) hydrogen bond with a backbone oxygen within the lipid bilayer. Hydrogen bonds are indicated by the dashed lines and their distances are shown in angstroms. (b) The side chains of the two lysine residues of this transmembrane helix are sufficiently long and flexible to form bonds with the head groups and water molecules of the polar face of the lipid bilayer. (c) The side chains of the two aspartic acid residues of this transmembrane helix can also reach the polar face of the bilayer but introduce distortion in the helix to do so. (d) The aromatic side chains of the two tyrosine residues of this transmembrane helix are oriented perpendicular to the axis of the membrane and parallel to the fatty acyl chains with which they interact.
310. Figure 4.20: Hydropathy plot for glycophorin A, a single membrane-spanning protein. Hydrophobicity is measured by the free energy required to transfer each segment of the polypeptide from a nonpolar solvent to an aqueous medium. Values above the 0 line are energy-requiring (+ΔGs), indicating they consist of stretches of amino acids that have predominantly nonpolar side chains. Peaks that project above the red-colored line are interpreted as a transmembrane domain.
311. Experimental Approaches to Identifying Conformational Changes within an Integral Membrane Protein
312. Figure 4.21: An experiment employing site-directed mutagenesis to learn about dynamic changes in the conformation of a membrane protein as it carries out its activity. The experimental strategy of the experiment and its results are discussed in the text. The cytoplasmic surface of the protein (lactose permease) is at the top. (a) The red spheres indicate the residues of the membrane protein that reacted with the alkylating agent NEM in the absence of a sugar to be transported. (b) The gold spheres denote the residues that are much more accessible to the alkylating agent when the protein is incubated with the sugar. The gold spheres in b are seen to cluster in a portion of the protein near the external medium (the periplasm in bacteria). The authors concluded that the gold spheres correspond to residues that line an outward-facing cavity, i.e., one that is open to the external medium. The results support a model of alternating access to the medium as depicted in part c. (Note the structure that is depicted in parts a–b is that of the inward-facing conformation as determined by X-ray crystallography. The structure of the outward-facing conformation has not been directly determined, which is why these types of labeling experiments remain important.)
313. Figure 4.22: Use of EPR spectroscopy to monitor changes in conformation of a bacterial K+ ion channel as it opens and closes. (a) EPR spectra from nitroxides that have been attached to cysteine residues near the cytoplasmic end of the four transmembrane helices that line the channel. The cysteine residue in each helix replaces a glycine residue that is normally at that position. The shapes of the spectra depend on the distances between unpaired electrons in the nitroxides on different subunits. (Nitroxides are described as “spin-labels,” and this technique is known as site-directed spin labeling.) (b) A highly schematic model of the ion channel in the open and closed states based on the data from part a. Opening of the channel is accompanied by the movement of the four nitroxide groups apart from one another.
314. Peripheral Membrane Proteins
315. Lipid-Anchored Membrane Proteins
316. REVIEW
317. 4.5: Membrane Lipids and Membrane Fluidity
318. Figure 4.23: The structure of the lipid bilayer depends on the temperature. The bilayer shown here is composed of two phospholipids: phosphatidylcholine and phosphatidylethanolamine. (a) Above the transition temperature, the lipid molecules and their hydrophobic tails are free to move in certain directions, even though they retain a considerable degree of order. (b) Below the transition temperature, the movement of the molecules is greatly restricted, and the entire bilayer can be described as a crystalline gel.
319. Table 4.2: Melting Points of the Common 18-Carbon Fatty Acids
320. The Importance of Membrane Fluidity
321. Maintaining Membrane Fluidity
322. Lipid Rafts
323. Figure 4.24: Lipid rafts. (a) Image of the upper surface of an artificial lipid bilayer containing phosphatidylcholine, which appears as the black background, and sphingomyelin molecules, which organize themselves spontaneously into the orange-colored rafts. The yellow peaks show the positions of a GPI-anchored protein, which is almost exclusively raft-associated. This image is provided by an atomic force microscope, which measures the height of various parts of the specimen at the molecular level. (b) Schematic model of a lipid raft within a cell. The outer leaflet of the raft consists primarily of cholesterol (yellow) and sphingolipids (red head groups). Phosphatidylcholine molecules (blue head groups) with long saturated fatty acids also tend to concentrate in this region. GPI-anchored proteins are thought to become concentrated in lipid rafts. The lipids in the outer leaflet of the raft have an organizing effect on the lipids of the inner leaflet. As a result, the inner-leaflet raft lipids consist primarily of cholesterol and glycerophospholipids with long saturated fatty acyl tails. The inner leaflet tends to concentrate lipid-anchored proteins, such as Src kinase, that are involved in cell signaling. (The controversy over the existence of lipid rafts is discussed in Nature Revs. Mol. Cell Biol. 11:688, 2010 and Science 334:1046, 2011.)
324. REVIEW
325. 4.6: The Dynamic Nature of the Plasma Membrane
326. Figure 4.25: The possible movements of phospholipids in a membrane. The types of movements in which membrane phospholipids can engage and the approximate time scales over which they occur. Whereas phospholipids move from one leaflet to another at a very slow rate, they diffuse laterally within a leaflet rapidly. Lipids lacking polar groups, such as cholesterol, can move across the bilayer quite rapidly.
327. The Diffusion of Membrane Proteins after Cell Fusion
328. Figure 4.26: The use of cell fusion to reveal mobility of membrane proteins. (a) Outline of the experiment in which human and mouse cells were fused (steps 1–2) and the distribution of the proteins on the surface of each cell were followed in the hybrids over time (steps 3–4). Mouse membrane proteins are indicated by solid circles, human membrane proteins by open circles. Locations of human and mouse proteins in the plasma membranes of the hybrid cells were monitored by interaction with fluorescent red and fluorescent green antibodies, respectively. (b) Micrograph showing a fused cell in which mouse and human proteins are still in their respective hemispheres (equivalent to the hybrid in step 3 of part a).
329. Restrictions on Protein and Lipid Mobility
330. Figure 4.27: Measuring the diffusion rates of membrane proteins by fluorescence recovery after photobleaching (FRAP)
331. Figure 4.28: Patterns of movement of integral membrane proteins.
332. Control of Membrane Protein Mobility
333. Membrane Lipid Mobility
334. Figure 4.29: Experimental demonstration that diffusion of phospholipids within the plasma membrane is confined. (a) The track of a single labeled unsaturated phospholipid is followed for 56 ms as it diffuses within the plasma membrane of a rat fibroblast. Phospholipids diffuse freely within a confined compartment before hopping into a neighboring compartment. The rate of diffusion within a compartment is as rapid as that expected by unhindered Brownian movement. However, the overall rate of diffusion of the phospholipid appears slowed because the molecule must hop a barrier to continue its movement. The movement of the phospholipid within each compartment is represented by a single color. (b) The same experiment shown in a is carried out for 33 ms in an artificial bilayer, which lacks the “picket fences” present in a cellular membrane. The much more open, extended trajectory of the phospholipid can now be explained by simple, unconfined Brownian movement. For the sake of comparison, fake compartments were assigned in b and indicated by different colors.
335. Membrane Domains and Cell Polarity
336. Figure 4.30: Differentiated functions of the plasma membrane of an epithelial cell. The apical surface of this intestinal epithelial cell contains integral proteins that function in ion transport and hydrolysis of disaccharides, such as sucrose and lactose; the lateral surface contains integral proteins that function in intercellular interaction; and the basal surface contains integral proteins that function in the association of the cell with the underlying basement membrane.
337. Figure 4.31: Differentiation of the mammalian sperm plasma membrane as revealed by fluorescent antibodies. (a–c) Three pairs of micrographs, each showing the distribution of a particular protein at the cell surface as revealed by a bound fluorescent antibody. The three proteins are localized in different parts of the continuous sperm membrane. Each pair of photographs shows the fluoresence pattern of the bound antibody and a phase contrast micrograph of the same cell. (d) Diagram summarizing the distribution of the proteins.
338. The Red Blood Cell: An Example of Plasma Membrane Structure
339. Integral Proteins of the Erythrocyte Membrane
340. Figure 4.32: The plasma membrane of the human erythrocyte. (a) Scanning electron micrograph of human erythrocytes. (b) Micrograph showing plasma membrane ghosts, which were isolated by allowing erythrocytes to swell and hemolyze as described in the text. (c) The results of SDS–polyacrylamide gel electrophoresis (SDS–PAGE) used to fractionate the proteins of the erythrocyte membrane, which are identified at the sides of the gel. (d) A model of the erythrocyte plasma membrane as viewed from the internal surface, showing the integral proteins embedded in the lipid bilayer and the arrangement of peripheral proteins that make up ere is simplified. The band 4.1 protein stabilthe membrane’s internal skeleton. The band 3 dimer shown hizes actin–spectrin complexes. (e) Electron micrograph showing the arrangement of the proteins of the inner membrane skeleton.
341. The Erythrocyte Membrane Skeleton
342. REVIEW
343. 4.7: The Movement of Substances Across Cell Membranes
344. The Energetics of Solute Movement
345. Figure 4.33: Four basic mechanisms by which solute molecules move across membranes. The relative sizes of the letters indicate the directions of the concentration gradients. (a) Simple diffusion through the bilayer, which always proceeds from high to low concentration. (b) Simple diffusion through an aqueous channel formed within an integral membrane protein or a cluster of such proteins. As in a, movement is always down a concentration gradient. (c) Facilitated diffusion in which solute molecules bind specifically to a membrane protein carrier (a facilitative transporter). As in a and b, movement is always from high to low concentration. (d) Active transport by means of a protein transporter with a specific binding site that undergoes a change in affinity driven with energy released by an exergonic process, such as ATP hydrolysis. Movement occurs against a concentration gradient. (e) Examples of each type of mechanism as it occurs in the membrane of an erythrocyte.
346. Diffusion of Substances through Membranes
347. The Diffusion of Water through Membranes
348. Figure 4.34: The relationship between partition coefficient and membrane permeability. In this case, measurements were made of the penetration of a variety of chemicals and drugs across the plasma membranes of the cells that line the capillaries of the brain. Substances penetrate by passage through the lipid bilayer of these cells. The partition coefficient is expressed as the ratio of solubility of a solute in octanol to its solubility in water. Permeability is expressed as penetrance (P) in cm/sec. For all but a few compounds, such as vinblastine and vincristine, penetrance is directly proportional to lipid solubility.
349. Figure 4.35: The effects of differences in the concentration of solutes on opposite sides of the plasma membrane. (a) A cell placed in a hypotonic solution (one having a lower solute concentration than the cell) swells because of a net gain of water by osmosis. (b) A cell in a hypertonic solution shrinks because of a net loss of water by osmosis. (c) A cell placed in an isotonic solution maintains a constant volume because the inward flux of water is equal to the outward flux.
350. Figure 4.36: The effects of osmosis on a plant cell. (a) Aquatic plants living in freshwater are surrounded by a hypotonic environment. Water therefore tends to flow into the cells, creating turgor pressure. (b) If the plant is placed in a hypertonic solution, such as seawater, the cell loses water, and the plasma membrane pulls away from the cell wall. (Ed Reschke.)
351. Figure 4.37: Passage of water molecules through an aquaporin channel. (a) Snapshot from a molecular dynamics simulation of a stream of water molecules (red and white spheres) passing in single file through the channel in one of the subunits of an aquaporin molecule residing within a membrane. (b) A model describing the mechanism by which water molecules pass through an aquaporin channel with the simultaneous exclusion of protons. Nine water molecules are shown to be lined up in single file along the wall of the channel. Each water molecule is depicted as a red circular O atom with two associated Hs. In this model, the four water molecules at the top and bottom of the channel are oriented, as the result of their interaction with the carbonyl (C=O) groups of the protein backbone (page 51), with their H atoms pointed away from the center of the channel. These water molecules are able to form hydrogen bonds (dashed lines) with their neighbors. In contrast, the single water molecule in the center of the channel is oriented in a position that prevents it from forming hydrogen bonds with other water molecules, which has the effect of interrupting the flow of protons through the channel. Animations of aquaporin channels can be found at www.nobelprize.org/nobel_prizes/chemistry/laureates/2003/animations.html
352. The Diffusion of Ions through Membranes
353. Figure 4.38: Measuring ion channel conductance by patch-clamp recording. (a) In this technique, a highly polished glass micropipette is placed against a portion of the outer surface of a cell, and suction is applied to seal the rim of the pipette against the plasma membrane. Because the pipette is wired as an electrode (a microelectrode), a voltage can be applied across the patch of membrane enclosed by the pipette, and the responding flow of ions through the membrane channels can be measured. As indicated in the figure, the micropipette can enclose a patch of membrane containing a single ion channel, which allows investigators to monitor the opening and closing of a single gated channel, as well as its conductance at different applied voltages. (b) The micrograph shows patch-clamp recordings being made from a single photoreceptor cell of the retina of a salamander. One portion of the cell is drawn into a glass micropipette by suction, while a second micropipette-electrode (lower right) is sealed against a small patch of the plasma membrane on another portion of the cell.
354. Figure 4.39: Three-dimensional structure of the bacterial KcsA channel and the selection of K+ ions. This K+ ion channel consists of four subunits, two of which are shown here. Each subunit is comprised of M1 and M2 helices joined by a P (pore) segment consisting of a short helix and a nonhelical portion that lines the channel through which the ions pass. A portion of each P segment contains a conserved pentapeptide (GYGVT) whose residues line the selectivity filter that screens for K+ ions. The oxygen atoms of the carbonyl groups of these residues project into the channel where they can interact selectively with K+ ions (indicated by the red mesh objects) within the filter. As indicated in the top inset, the selectivity filter contains four rings of carbonyl O atoms and one ring of threonyl O atoms; each of these five rings contains four O atoms, one donated by each subunit. The diameter of the rings is just large enough so that eight O atoms can coordinate a single K+ ion, replacing its normal water of hydration. Although four K+ binding sites are shown, only two are occupied at one time.
355. Figure 4.40: Schematic illustration of the hinge-bending model for the opening of the bacterial KcsA channel. The M2 helices from each subunit bend outward at a specific glycine residue, which opens the gate at the intracellular end of the channel to K+ ions.
356. Figure 4.41: The structure of one subunit of a eukaryotic, voltage-gated K+ channel. A two-dimensional portrait of a K+ channel subunit showing its six transmembrane helices and a portion of the polypeptide (called the pore helix or P) that dips into the protein to form part of the channel’s wall. The inset shows the sequence of amino acids of the positively charged S4 helix of the Drosophila K+ Shaker ion channel, which serves as a voltage sensor. The positively charged side chains are situated at every third residue along the otherwise hydrophobic helix. This member of the Kv family is called a Shaker channel because flies with certain mutations in the protein shake vigorously when anesthesized with ether. The Shaker channel was the first K+ channel to be identified and cloned in 1987.
357. Figure 4.42: Three-dimensional structure of a voltage-gated mammalian K+ channel. (a) The crystal structure of the entire tetrameric Kv1.2 channel, a member of the Shaker family of K+ ion channels found in nerve cells of the brain. The transmembrane portion is shown in red, and the cytoplasmic portion in blue. The potassium ion binding sites are indicated in green. (b) Ribbon drawing of the same channel shown in a, with the four subunits that make up the channel shown in different colors. If you focus on the red subunit, you can see (1) the spatial separation between the voltage-sensing and pore domains of the subunit and (2) the manner in which the voltage-sensing domain from each subunit is present on the outer edge of the pore domain of a neighboring subunit. The cytoplasmic portion of this particular channel consists of a T1 domain, which is part of the channel polypeptide itself, and a separate β polypeptide. (c) Ribbon drawing of a single subunit showing the spatial orientation of the six membrane-spanning helices (S1–S6) and also the presence of the S4–S5 linker helix, which connects the voltage-sensing and pore domains. This linker transmits the signal from the S4 voltage sensor that opens the channel. The inner surface of the channel below the pore domain is lined by the S6 helix (roughly similar to the M2 helix of the bacterial channel shown in Figure 4.39). The channel shown here is present in the open configuration with the S6 helices curved outward (compare to Figure 4.40) at the site marked PVP (standing for Pro-Val-Pro, which is likely the amino acid sequence of the “hinge”).
358. Figure 4.43: Conformational states of a voltage-gated K+ ion channel. (a) Three-dimensional model of a eukaryotic K+ ion channel. Inactivation of channel activity occurs as one of the inactivation peptides, which dangle from the cytoplasmic portion of the complex, fits into the cytoplasmic opening of the channel. (b) Schematic representation of a view into a K+ ion channel, perpendicular to the membrane from the cytoplasmic side, showing the channel in the closed (resting), open, and inactivated state.
359. Facilitated Diffusion
360. Figure 4.44: Facilitated diffusion. A schematic model for the facilitated diffusion of glucose depicts the alternating conformation of a carrier that exposes the glucose binding site to either the inside or outside of the membrane.
361. Figure 4.45: The kinetics of facilitated diffusion as compared to that of simple physical diffusion.
362. The Glucose Transporter: An Example of Facilitated Diffusion
363. Active Transport
364. Primary Active Transport: Coupling Transport to ATP Hydrolysis
365. Table 4.3: Ion Concentrations Inside and Outside of a Typical Mammalian Cell
366. FIGURE IN FOCUS
367. Figure 4.46: The Na+/K+-ATPase. (a) Simplified schematic model of the transport cycle as described in the text. Note that the actual Na+/K+-ATPase is composed of at least two different membrane-spanning subunits: a larger α subunit, which carries out the transport activity, and a smaller β subunit, which functions primarily in the maturation and assembly of the pump within the membrane. A third (γ) subunit may also be present. (b) A model of the E2 conformation of the protein based on an X-ray crystallographic study. The cation binding sites are located deep within the transmembrane domain, which consists of 10 membrane-spanning helices. The two rubidium ions are located where the potassium ions would normally be bound.
368. Other Primary Ion Transport Systems
369. Figure 4.47: Control of acid secretion in the stomach. In the resting state, the H+/K+-ATPase molecules are present in the walls of cytoplasmic vesicles. Food entering the stomach triggers a cascade of hormone-stimulated reactions in the stomach wall leading to the release of histamine, which binds to a receptor on the surface of the acid-secreting parietal cells. Binding of histamine to its receptor stimulates a response that causes the H+/K+-ATPase-containing vesicles to fuse to the plasma membrane, forming deep folds, or canaliculi. Once at the surface, the transport protein is activated and pumps protons into the stomach cavity against a concentration gradient (indicated by the size of the letters). The heartburn drugs Prilosec, Nexium, and Prevacid block acid secretion by directly inhibiting the H+/K+-ATPase, whereas several other acid-blocking medications interfere with activation of the parietal cells. Acid-neutralizing medications provide basic anions that combine with the secreted protons.
370. Using Light Energy to Actively Transport Ions
371. Figure 4.48: Bacteriorhodopsin: a light-driven proton pump. The protein contains seven membrane-spanning helices and a centrally located retinal group (shown in purple), which serves as the light-absorbing element (chromophore). Absorption of a photon of light causes a change in the electronic structure of retinal, leading to the transfer of a proton from the —NH+ group to a closely associated, negatively charged aspartic acid residue (#85) (step 1). The proton is then released to the extracellular side of the membrane (step 2) by a relay system consisting of several amino acid residues (Asp82, Glu204, and Glu194). The spaces between these residues are filled with hydrogen-bonded water molecules that help shuttle protons along the pathway. The deprotonated retinal is returned to its original state (step 3) when it accepts a proton from an undissociated aspartic acid residue (Asp96) located near the cytoplasmic side of the membrane. Asp96 is then reprotonated by a H+ from the cytoplasm (step 4). Asp85 is deprotonated (step 5) prior to receiving a proton from retinal in the next pumping cycle. As a result of these events, protons move from the cytoplasm to the cell exterior through a central channel in the protein.
372. Secondary Active Transport (or Cotransport): Coupling Transport to Existing Ion Gradients
373. Figure 4.49: Secondary transport: the use of energy stored in an ionic gradient. The Na+/K+-ATPase residing in the plasma membrane of the lateral surface maintains a very low cytosolic concentration of Na+. The Na+ gradient across the plasma membrane represents a storage of energy that can be tapped to accomplish work, such as the transport of glucose by a Na+/glucose cotransporter located in the apical plasma membrane. Once transported across the apical surface into the cell, the glucose molecules diffuse to the basal surface where they are carried by a glucose facilitative transporter out of the cell and into the bloodstream. The relative size of the letters indicates the directions of the respective concentration gradients. Two Na+ ions are transported for each glucose molecule; the 2;1 Na+/glucose provides a much greater driving force for moving glucose into the cell than a 1;1 ratio.
374. Figure 4.50: A schematic model of the transport cycle of a secondary transporter. Four different conformational states during the transport cycle of a bacterial symporter of the LeuT family are shown. The protein actively transports the amino acid leucine into the cell using the established Na+ ion gradient as its source of energy. In step 1, the outer gate in the protein is open, whch allows both Na+ and leucine to reach their binding sites from the extracellular space. In step, 2, the outer gate closes, occluding the substrates within the protein. In step 3, a second leucine molecule binds to another site just outside the outer gate. In step 4, the inner gate opens and the substrates are released into the cytoplasm. The protein returns to its original state when the inner gate is closed and the outer gate is opened.
375. THE HUMAN PERSPECTIVE: Defects in Ion Channels and Transporters as a Cause of Inherited Disease
376. Table 1
377. Figure 1: An explanation for the debilitating effects on lung function from the absence of the CFTR protein. In the airway epithelium of a normal individual, water flows out of the epithelial cells in response to the outward movement of ions, thus hydrating the surface mucous layer. The hydrated mucous layer, with its trapped bacteria, is readily moved out of the airways. In the airway epithelium of a person with cystic fibrosis, the abnormal movement of ions causes water to flow in the opposite direction, thus dehydrating the mucous layer. As a result, trapped bacteria cannot be moved out of the airways, which allows them to proliferate as a biofilm (page 13) and cause chronic infections.
378. REVIEW
379. 4.8: Membrane Potentials and Nerve Impulses
380. Figure 4.51: The structure of a nerve cell. (a) Schematic drawing of a simple neuron with a myelinated axon. As the inset shows, the myelin sheath comprises individual Schwann cells that have wrapped themselves around the axon. The sites where the axon lacks myelin wrapping are called nodes of Ranvier. (Note: Myelin-forming cells within the central nervous system are called oligodendrocytes rather than Schwann cells.) (b) A composite micrograph of a single rat hippocampal neuron with cell body and dendrites (purple) and an axon 1 cm in length (red). Motor nerve cells in larger mammals can be 100 times this length.
381. The Resting Potential
382. Figure 4.52: Measuring a membrane’s resting potential.
383. The Action Potential
384. Figure 4.53: Formation of an action potential. (a) Time 1, upper left box: The membrane in this region of the nerve cell exhibits the resting potential, in which only the K+ leak channels are open and the membrane voltage is approximately −70 mV. Time 2, upper middle box, shows the depolarization phase: The membrane has depolarized beyond the threshold value, opening the voltage-regulated sodium gates, leading to an influx of Na+ ions (indicated in the permeability change in the lower graph). The increased Na+ permeability causes the membrane voltage to temporarily reverse itself, reaching a value of approximately +40 mV in the squid giant axon (time 2). It is this reversal of membrane potential that constitutes the action potential. Time 3, upper right box, shows the repolarization phase: Within a tiny fraction of a second, the sodium gates are inactivated and the potassium gates open, allowing potassium ions to diffuse across the membrane (lower part of the drawing) and establish an even more negative potential at that location (−80 mV) than that of the resting potential. Almost as soon as they open, the potassium gates close, leaving the potassium leak channels as the primary path of ion movement across the membrane and reestablishing the resting potential. (b) A summary of the voltage changes that occur during an action potential, as described in part a.
385. Propagation of Action Potentials as an Impulse
386. Figure 4.54: Propagation of an impulse results from the local flow of ions. An action potential at one site on the membrane depolarizes an adjacent region of the membrane, triggering an action potential at the second site. The action potential can only flow in the forward direction because the portion of the membrane that has just experienced an action potential remains in a refractory period.
387. Speed Is of the Essence
388. Figure 4.55: Saltatory conduction. During saltatory conduction, only the membrane in the nodal region of the axon becomes depolarized and capable of forming an action potential. This is accomplished as current flows directly from an activated node to the next resting node along the axon.
389. Figure 4.56: The neuromuscular junction is the site where branches from a motor axon form synapses with the muscle fibers of a skeletal muscle. The left inset shows the synaptic vesicles residing within the terminal knob of the axon and the narrow synaptic cleft between the terminal knob and the postsynaptic target cell. The right inset shows the terminal knob pressed closely against the muscle cell plasma membrane. Neurotransmitter molecules (red) released from synaptic vesicles of the presynaptic neuron are binding to receptors (yellow) on the surface of the muscle cell (blue).
390. Neurotransmission: Jumping the Synaptic Cleft
391. Figure 4.57: The sequence of events during synaptic transmission with acetylcholine as the neurotransmitter. During steps 1–4, a nerve impulse reaches the terminal knob of the axon, calcium gates open leading to an influx of Ca2+, and acetylcholine is released from synaptic vesicles and binds to receptors on the postsynaptic membrane. If the binding of the neurotransmitter molecules causes a depolarization of the postsynaptic membrane (as in 5a), a nerve impulse may be generated there (6). If, however, the binding of neurotransmitter causes a hyperpolarization of the postsynaptic membrane (5b), the target cell is inhibited, making it more difficult for an impulse to be generated in the target cell by other excitatory stimulation. The breakdown of the neurotransmitter by acetylcholinesterase is not shown.
392. Actions of Drugs on Synapses
393. Synaptic Plasticity
394. REVIEW
395. EXPERIMENTAL PATHWAYS: The Acetylcholine Receptor
396. Figure 1 The electric organs of Torpedo consist of stacks of modified neuromuscular junctions located on each side of the body.
397. Figure 2 Steps used in the isolation of the nAChR. (a) Structure of a synthetic compound, CT5263, that was attached to sepharose beads to form an affinity column. The ends of the compound projecting from the beads resemble acetylcholine, causing both acetylcholinesterase (AChE) and the nicotinic acetylcholine receptor (nAChR) to bind to the beads. (b) When the Triton X-100 extract was passed through the column, both of the acetylcholine-binding proteins stuck to the beads, while the remaining dissolved protein (about 90 percent of the total protein in the extract) passed directly through the column. Subsequent passage of a solution of 10−3 M flaxedil through the column released the bound nAChR, without disturbing the bound AChE (which was subsequently eluted with 1 M NaCl).
398. Figure 3 The top portion of the figure shows an SDS–polyacrylamide gel following electrophoresis of a preparation of the purified nAChR. The receptor consists of four different subunits whose molecular weights are indicated. Prior to electrophoresis, the purified receptor preparation was incubated with a radioactive compound (3H−MBTA) that resembles acetylcholine and binds to the acetylcholine-binding site of the nAChR. Following electrophoresis, the gel was sliced into 1-mm sections and the radioactivity of each slice determined. All of the radioactivity was bound to the 39,000 dalton subunit, indicating this subunit contains the ACh-binding site. The dotted line indicates the light absorbance of each fraction, which provides a measure of the total amount of protein present in that fraction. The heights of the peaks provide a measure of the relative amounts of each of the subunits in the protein. All of the subunits are present in equal numbers except the smallest subunit (the α subunit, which contains the ACh-binding site), which is present in twice the number of copies.
399. Figure 4 Electron micrograph of negatively stained, receptor-rich membranes from the electric organ of an electric fish showing the dense array of nAChR molecules. Each receptor molecule is seen as a small whitish circle with a tiny black dot in its center; the dot corresponds to the central channel, which has collected electron-dense stain.
400. Figure 5 (a) An electron density map of a slice through the nAChR obtained by analyzing electron micrographs of tubular crystals of Torpedo membranes embedded in ice. These analyses have allowed researchers to reconstruct the three-dimensional appearance of a single nAChR protein as it resides within the membrane. The continuous contours indicate lines of similar density greater than that of water. The two dark, bar-shaped lines represent the a helices that line the channel at its narrowest point. (b) Schematic diagram of the nAChR showing the arrangement of the subunits and a cross-sectional representation of the protein. Each of the five subunits contains four membrane-spanning helices (M1–M4). Of these, only the inner helix (M2) lines the pore, and is the subject of the remainder of the discussion.
401. Figure 6: Ribbon drawings illustrating the proposed changes that occur within the nAChR upon binding of acetylcholine. Only the two alpha subunits of the receptor are shown. In the closed state (left) the pore is blocked (pink patch) by the close apposition of a ring of hydrophobic residues (the valine and leucine side chains of these residues on the alpha subunits are indicated by the small ball-and-stick models at the site of pore constriction). The diameter of the pore at its narrowest point is about 6 Å, which is not sufficient for a hydrated Na+ ion to pass. Although it is wide enough for passage of a dehydrated Na+ ion, the wall of the channel lacks the polar groups that would be required to substitute for the displaced shell of water molecules (as occurs in the selectivity filter of the K+ channel, Figure 4.39). Following binding of ligand, a conformational change is proposed, which leads to a small rotation of the inner β sheets in the ligand-binding domain of the alpha subunits (curved arrows on left figure). This, in turn, induces a rotational movement of the inner transmembrane helices of the subunits, expanding the diameter of the pore, which allows the flow of Na+ ions through the open state of the channel (right). The relevant moving parts are shown in blue.
402. References
403. Synopsis
404. Analytic Questions
405. 5: Aerobic Respiration and the Mitochondrion
406. Micrograph of a mammalian fibroblast that has been fixed and stained with fluorescent antibodies that reveal the distribution of the mitochondria (green) and the microtubules of the cytoskeleton (red). The mitochondria are seen as an extensive network or reticulum that extends through much of the cell.
407. 5.1: Mitochondrial Structure and Function
408. Figure 5.1: Mitochondria.
409. Figure 5.2: Mitochondrial fusion and fission.
410. Mitochondrial Membranes
411. FIGURE IN FOCUS
412. Figure 5.3: The structure of a mitochondrion.
413. The Mitochondrial Matrix
414. Figure 5.4: Porins.
415. Figure 5.5: An overview of carbohydrate metabolism in eukaryotic cells.
416. REVIEW
417. 5.2: Oxidative Metabolism in the Mitochondrion
418. Figure 5.6: An overview of glycolysis showing some of the key steps.
419. The Tricarboxylic Acid (TCA) Cycle
420. Figure 5.7: The tricarboxylic acid (TCA) cycle,
421. Figure 5.8: Catabolic pathways generate compounds that are fed into the TCA cycle.
422. The Importance of Reduced Coenzymes in the Formation of ATP
423. Figure 5.9: The glycerol phosphate shuttle.
424. Figure 5.10: A summary of the process of oxidative phosphorylation.
425. REVIEW
426. THE HUMAN PERSPECTIVE: The Role of Anaerobic and Aerobic Metabolism in Exercise
427. Figure 1: Skeletal muscles
428. 5.3: The Role of Mitochondria in the Formation of ATP
429. Oxidation–Reduction Potentials
430. Figure 5.11: Measuring the standard oxidation–reduction (redox) potential.
431. Table 5.1: Standard Redox Potentials of Selected Half-Reactions
432. Electron Transport
433. Types of Electron Carriers
434. Figure 5.12: Structures of the oxidized and reduced forms of three types of electron carriers.
435. Figure 5.13: Iron-sulfur centers.
436. Figure 5.14: The arrangement of several carriers in the electron-transport chain.
437. Figure 5.15: Experimental use of inhibitors to determine the sequence of carriers in the electron-transport chain.
438. Electron-Transport Complexes
439. Figure 5.16 Electron-tunneling pathways for the yeast cytochrome c-cytochrome c peroxidase complex.
440. Figure 5.17: The electron-transport chain of the inner mitochondrial membrane.
441. Figure 5.18: Experimental demonstration that cytochrome oxidase is a proton pump.
442. Figure 5.19: Structure and a proposed mechanism of action of complex I of the respiratory chain.
443. Complex I (or NADH dehydrogenase)
444. Complex II (or succinate dehydrogenase)
445. Complex III (or cytochrome bc1)
446. Complex IV (or cytochrome c oxidase)
447. A Closer Look at Cytochrome Oxidase
448. Figure 5.20: A summary of cytochrome oxidase activity.
449. REVIEW
450. 5.4: Translocation of Protons and the Establishment of a Proton-Motive Force
451. Figure 5.21: Visualizing the proton-motive force.
452. REVIEW
453. 5.5: The Machinery for ATP Formation
454. Figure 5.22: The machinery for ATP synthesis.
455. Figure 5.23: An experiment to drive ATP formation in membrane vesicles reconstituted with the Na+/K+-ATPase.
456. Figure 5.24: The structure of the bacterial ATP synthase.
457. The Structure of ATP Synthase
458. Figure 5.25: Visualizing the oligomeric c ring of a chloroplast ATP synthase.
459. The Basis of ATP Formation According to the Binding Change Mechanism
460. Evidence to Support the Binding Change Mechanism and Rotary Catalysis
461. Figure 5.26: The structural basis of catalytic site conformation.
462. Figure 5.27: The binding change mechanism for ATP synthesis.
463. Figure 5.28: Direct observation of rotational catalysis.
464. Using the Proton Gradient to Drive the Catalytic Machinery: The Role of the F0 Portion of ATP Synthase
465. Figure 5.29: A model in which proton diffusion is coupled to the rotation of the c ring of the F0 complex.
466. Figure 5.30: Summary of the major activities during aerobic respiration in a mitochondrion.
467. Other Roles for the Proton-Motive Force in Addition to ATP Synthesis
468. REVIEW
469. 5.6: Peroxisomes
470. Figure 5.31: The structure and function of peroxisomes.
471. REVIEW
472. Figure 5.32: Glyoxysome localization within plant seedlings.
473. THE HUMAN PERSPECTIVE: Diseases that Result from Abnormal Mitochondrial or Peroxisomal Function
474. Mitochondria
475. Figure 1: Mitochondrial abnormalities in skeletal muscle.
476. Figure 2: A premature-aging phenotype caused by increased mutations in mtDNA.
477. Peroxisomes
478. Synopsis
479. Analytic Questions
480. 6: Photosynthesis and the Chloroplast
481. Light micrograph of a living leaf cell from Arabidopsis thaliana. The cell contains a number of chloroplasts—the organelles that house the plant’s photosynthetic machinery. Chloroplasts normally divide by binary fission in which a single constriction splits the organelle into two equal daughters. This cell is from a mutant plant that is characterized by asymmetric fission. The highly elongated chloroplast has initiated fission asymmetrically at several sites, as indicated by the multiple constrictions. Mutants have proven invaluable in the identification of genes involved in all types of cellular processes. It is only when a gene malfunctions that its effects usually become visible, providing researchers with an idea of the gene’s normal function—in this case a role in chloroplast fission.
482. Figure 6.1: Photosynthetic green sulfur bacteria are present as a ring of peripheral cells that are living in a symbiotic relationship with a single anaerobic, heterotrophic bacterium in the center of the “colony.” The heterotrophic bacterium receives organic matter produced by the photosynthetic symbionts. Photosynthetic vesicles containing the light-capturing machinery are visible in the green sulfur bacteria.
483. 6.1: Chloroplast Structure and Function
484. Figure 6.2: The functional organization of a leaf. The section of the leaf shows several layers of cells that contain chloroplasts distributed within their cytoplasm. These chloroplasts carry out photosynthesis, providing raw materials and chemical energy for the entire plant.
485. Figure 6.3: The internal structure of a chloroplast. (a) Transmission electron micrograph through a single chloroplast. The internal membrane is arranged in stacks (grana) of disk-like thylakoids that are physically separate from the outer double membrane that forms the envelope. (b) Schematic diagram of a chloroplast showing the outer double membrane and the thylakoid membranes.
486. REVIEW
487. Figure 6.4: Thylakoid membranes. Electron micrograph of a section through a portion of a chloroplast showing the stacked grana thylakoids, which are connected to one another by unstacked stroma thylakoids (or stroma lamellae). The dark spheres are osmium-stained lipid granules.
488. 6.2: An Overview of Photosynthetic Metabolism
489. Figure 6.5: An overview of the energetics of photosynthesis and aerobic respiration.
490. REVIEW
491. 6.3: The Absorption of Light
492. Photosynthetic Pigments
493. Figure 6.6: The structure of chlorophyll a. The molecule consists of a porphyrin ring (which in turn is constructed of four smaller pyrrole rings) with a magnesium ion at its center and a long hydrocarbon tail. The green shading around the edge of the porphyrin indicates the delocalization of electrons that form a cloud. The structure of the magnesium-containing porphyrin of chlorophyll can be compared to the iron-containing porphyrin of a heme shown in Figure 5.12. Chlorophyll b and bacteriochlorophyll a contain specific substitutions as indicated. For example, the —CH3 group on ring II is replaced by a —CHO group in chlorophyll b. Chlorophyll a is present in all oxygen-producing photosynthetic organisms, but it is absent in the various sulfur bacteria. In addition to chlorophyll a, chlorophyll b is present in all higher plants and green algae. Others not shown are chlorophyll c, present in brown algae, diatoms, and certain protozoa, and chlorophyll d, found in red algae. Bacteriochlorophyll is found only in green and purple bacteria, organisms that do not produce O2 during photosynthesis.
494. Figure 6.7: Absorption spectrum for several photosynthetic pigments of higher plants. The background shows the colors that we perceive for the wavelengths of the visible spectrum. Chlorophylls absorb most strongly in the violet, blue, and red regions of the spectrum, while carotenoids (e.g., β-carotene) also absorb into the green region. Red algae and cyanobacteria contain additional pigments (phycobilins) that absorb in the middle bands of the spectrum.
495. Figure 6.8: Action spectrum for photosynthesis. The action spectrum (red-colored line) indicates the relative efficiency with which light of various wavelengths is able to promote photosynthesis in the leaves of a plant. An action spectrum can be generated by measuring the O2 produced by the leaves following exposure to various wavelengths. The black lines indicate the absorption spectra of each of the major photosynthetic pigments. The green line shows the combined absorption spectrum of all pigments.
496. REVIEW
497. 6.4: Photosynthetic Units and Reaction Centers
498. Figure 6.9: The transfer of excitation energy. Energy is transferred randomly through a network of pigment molecules that absorb light of increasingly longer wavelength until the energy reaches a reaction-center chlorophyll, which transfers an excited electron to a primary acceptor, as described later in the chapter.
499. Oxygen Formation: Coordinating the Action of Two Different Photosynthetic Systems
500. Figure 6.10: An overview of the flow of electrons during the light-dependent reactions of photosynthesis. The events depicted in this schematic drawing are described in detail in the following pages. The energy content of electrons is given in volts. To convert these values to calories, multiply by the Faraday constant, 23.06 kcal/V. For example, a difference of 2.0 V corresponds to an energy difference of 46 kcal/mol of electrons. This can be compared to the energy of red light (680 nm), which contains about 42 kcal/mol of photons.
501. PSII Operations: Obtaining Electrons by Splitting Water
502. Figure 6.11: The functional organization of photosystem II. (a) A schematic model of the huge protein–pigment complex, which catalyzes the light-driven oxidation of water and reduction of plastoquinone. The path taken by electrons through PSII is indicated by the yellow arrows. Events begin with the absorption of light by an antenna pigment in the outer light-harvesting complex (LHCII). Energy is transferred from LHCII through an inner antenna pigment–protein complex to a P680 reaction-center chlorophyll a, which is one of four closely spaced chlorophyll a molecules (the P680 dimer and two accessory chlorophyll a molecules). Absorption of this energy by P680 excites an electron, which is transferred to pheophytin (Pheo) (step 1), the primary electron acceptor of PSII. (Pheophytin is a chlorophyll molecule that lacks the Mg2+ ion.) The electron subsequently passes to a plastoquinone PQA (step 2) and then through a nonheme Fe2+ to PQB (step 3) to form a negatively charged free radical PQB.−. Absorption of a second photon sends a second electron along the same pathway, converting the acceptor to PQB2− (step 4). Two protons then enter from the stroma (step 5) generating PQH2, which is released into the lipid bilayer and replaced by a new oxidized PQB molecule (step 6). As the above events are occurring, electrons are moving from H2O by way of Tyrz to the positively charged reaction-center pigment (steps B and A). Thus, overall, PSII catalyzes the transfer of electrons from water to plastoquinone. The oxidation of two molecules of H2O to release one molecule of O2 generates two molecules of PQH2. Because the oxidation of water releases protons into the thylakoid lumen and the reduction of PQB2− removes protons from the stroma, the operation of PSII makes a major contribution to formation of a H+ gradient. The figure shows one monomer of a dimeric PSII complex. (b) The oxygen-evolving complex contains a Mn4CaO5 cluster whose structure has been determined to a resolution of 1.9 Å by X-ray crystallography. Three Mn atoms (#1-3), a Ca atom, and four O atoms (#1, 2, 3, 5) are arranged as part of an asymmetric cube-like cluster with a bridge to a fourth Mn and fifth O atom at nearby sites. Two water molecules are bound to the Ca and two more are bound to Mn4. It is likely that reaction between the oxygen atoms of two of these bound water molecules leads to formation of an O = O bond.
503. The Flow of Electrons from PSII to Plastoquinone
504. Figure 6.12: Plastoquinone. The acceptance of two electrons and two protons reduces PQ (plastoquinone) to PQH2 (plastoquinol). The intermediates are similar to those shown in Figure 6.13c for ubiquinone of the mitochondrion.
505. The Flow of Electrons from Water to PSII
506. Figure 6.13: Measuring the kinetics of O2 release. The plot shows the response by isolated chloroplasts that have been kept in the dark to a succession of light flashes of very short duration. The amount of oxygen released peaks with every fourth flash of light. The first peak occurs after three flashes (rather than four) because most of the manganese cluster is present in the S1 state (one oxidizing equivalent) when kept in the dark. The oscillations become damped as the number of flashes increases.
507. From PSII to PSI
508. Figure 6.14: Electron transport between PSII and PSI. The flow of a pair of electrons is indicated by the yellow arrow. Cytochrome b6f operates in a manner very similar to that of cytochrome bc1 in the mitochondria and engages in a Q cycle (not discussed in the text) that translocates four protons for every pair of electrons that moves through the complex. PQH2 and PC (plastocyanin) are mobile carriers that can transport electrons between distantly separated photosystems.
509. PSI Operations: The Production of NADPH
510. Figure 6.15: The functional organization of photosystem I. The path taken by electrons is indicated by the yellow arrow. Events begin with the absorption of light by an antenna pigment and transfer of the energy to a P700 chlorophyll at the PSI reaction center. Absorption of energy by P700 causes the excitation of an electron and its transfer (step 1) to A0, which is the primary electron acceptor of PSI. The electron subsequently passes to A1 (step 2) and then to an iron-sulfur center named FX (step 3). From FX the electron is transferred (step 4) through two more iron-sulfur centers (FA and FB), which are bound by a peripheral protein at the stromal side of the membrane. The electron is finally transferred to ferredoxin, a small iron-sulfur protein (step 5) that is external to the PSI complex. When two different ferredoxin molecules have accepted an electron, they act together to reduce a molecule of NADP+ to NADPH (step 6). The electron-deficient reaction-center pigment (P700+) is reduced by an electron donated by plastocyanin (step A).
511. An Overview of Photosynthetic Electron Transport
512. FIGURE IN FOCUS
513. Figure 6.16: Summary of the light-dependent reactions. (a) Three-dimensional structures of the proteins of the thylakoid membrane that carry out the light-dependent reactions of photosynthesis. Of the four major protein complexes, PSII and cytochrome b6f are present in the membrane as dimers, whereas PSI and the ATP synthase (shown in greater detail in Figure 5.24) are present as monomers. (b) Summary of the flow of electrons from H2O to NADPH through the three transmembrane complexes. This figure shows the estimated number of protons translocated through the membrane as the result of the oxidation of two molecules of water, yielding two pairs of electrons. The ATP synthase of the thylakoid membranes is also shown (see Section 5.5 for discussion of the ATP-synthesizing enzyme). Approximately four protons are required for the synthesis of each molecule of ATP (page 205).
514. Killing Weeds by Inhibiting Electron Transport
515. REVIEW
516. 6.5: Photophosphorylation
517. Noncyclic Versus Cyclic Photophosphorylation
518. Figure 6.17: Simplified scheme for cyclic photophosphorylation. Absorption of light by PSI excites an electron, which is transferred to ferredoxin (step 1) and on to cytochrome b6f (step 2), plastocyanin (step 3), and back to P700+ (step 4). In the process, protons are translocated by cytochrome b6f to form a gradient utilized for ATP synthesis (step 5). Another cyclic pathway for electron transport that involves movement of electrons from PSI through NADPH to cytochrome b6f is not shown.
519. REVIEW
520. 6.6: Carbon Dioxide Fixation and the Synthesis of Carbohydrate
521. Carbohydrate Synthesis in C3 Plants
522. Figure 6.18: Chromatogram showing the results of an experiment in which algal cells were incubated for 5 seconds in [14C]O2 prior to immersion in alcohol. One spot, which corresponds to 3-phosphoglycerate (labeled PGA), contains most of the radioactivity.
523. Figure 6.19: Converting CO2 into carbohydrate. (a) The reaction catalyzed by ribulose bisphosphate carboxylase oxygenase (Rubisco) in which CO2 is fixed by linkage to RuBP. The product rapidly splits into two molecules of 3-phosphoglycerate (PGA). (b) An abbreviated version of the Calvin cycle showing the fate of 6 molecules of CO2 that are fixed by combination with 6 molecules of RuBP. (Numerous reactions have been deleted). The fixation of CO2 is indicated in step 1. In step 2, the 12 PGA molecules are phosphorylated via ATP hydrolysis to form 12 1,3-bisphosphoglycerate (BPG) molecules, which are reduced in step 3 by electrons provided by NADPH to form 12 molecules of glyceraldehyde 3-phosphate (GAP). Here, 2 of the GAPs are drained away (step 4) to be used in the synthesis of sucrose in the cytosol, which can be considered the product of the light-independent reactions. The other 10 molecules are converted into 6 molecules of RuBP (step 5), which can act as the acceptor for 6 more molecules of CO2. The regeneration of 6 RuBPs requires the hydrolysis of 6 molecules of ATP. The NADPH and ATP used in the Calvin cycle represent the two high-energy products of the light-dependent reactions.
524. Figure 6.20: An overview of the various stages of photosynthesis.
525. Redox Control
526. Figure 6.21: Redox control of the Calvin cycle. In the light, ferredoxin is reduced, and a fraction of these electrons are transferred to the small protein thioredoxin, which reduces the disulfide groups of certain Calvin cycle enzymes, maintaining them in an active state. In the dark, electron flow to thioredoxin ceases, the sulfhydryl groups of the regulated enzymes become oxidized to the disulfide state, and the enzymes are inactivated.
527. Photorespiration
528. Figure 6.22: The reactions of photorespiration. Rubisco can catalyze two different reactions with RuBP as a substrate (shown in the enediol state within the plane; see Figure 6.19a). If the RuBP reacts with O2 (step 1b), the reaction produces an oxygenase intermediate (step 2b) that breaks down into 3-PGA and 2-phosphoglycolate (step 3b). The subsequent reactions of phosphoglycolate are shown in Figure 6.23. The eventual outcome of these reactions is the release of CO2, a molecule that the cell had previously expended energy to fix. In contrast, if the RuBP molecule reacts with CO2 (step 1a), the reaction produces a carboxylase intermediate (step 2a) that breaks down into two molecules of PGA (step 3a), which continue through the Calvin cycle.
529. Figure 6.23: The cellular basis of photorespiration. Electron micrograph of a portion of a leaf mesophyll cell of a tobacco plant showing a peroxisome (identified by its crystalline core) pressed against a pair of chloroplasts and close to a mitochondrion. The reactions of photorespiration that occur in each of these organelles are described in the text and shown superimposed on the organelles in which they occur. This series of reactions is referred to as the C2 cycle. The last steps of the cycle—the conversion of serine to glycerate in the peroxisome and then to 3-PGA in the chloroplast—are not shown.
530. Peroxisomes and Photorespiration
531. Carbohydrate Synthesis in C4 Plants
532. Figure 6.24: Structure and function in C4 plants. Electron micrograph of a transverse section through the leaf of a C4 plant showing the spatial relationship between the mesophyll and bundle-sheath cells. Superimposed on the micrograph are the reactions of CO2 fixation that occur in each type of cell. In step 1, CO2 is joined to PEP by the enzyme PEP carboxylase in a mesophyll cell that is located close to the leaf exterior. The four-carbon malate formed by the reaction is transported to the more centrally located bundle-sheath cell (step 2), where the CO2 is released. The CO2 becomes highly concentrated in the bundle-sheath cell, favoring the fixation of CO2 by Rubisco to form 3-PGA (step 3), which can be circulated through the Calvin cycle. The pyruvate formed when CO2 is released is shipped back to the mesophyll cell (step 4), where it is converted to PEP. Although the process requires the hydrolysis of ATP (step 5), the high CO2/O2 in the bundle-sheath cell minimizes the rate of photorespiration.
533. Carbohydrate Synthesis in CAM Plants
534. REVIEW
535. Synopsis
536. Analytic Questions
537. 7: Interactions Between Cells and Their Environment
538. Fluorescence micrograph of an endothelial cell—a type of cell that lines the inner surface of blood vessels. The cell is square because it has spread itself over a tiny square-shaped patch of an adhesive protein called fibronectin that was applied to a culture dish. The cell appears to be mounted in a green frame because it was treated with a green fluorescent antibody that binds to the cytoplasmic protein actin, a component of the cytoskeleton.
539. Figure 7.1: An overview of how cells are organized into tissues and how they interact with one another and with their extracellular environment. In this schematic diagram of a section through human skin, the cells of the epidermis are seen to adhere to one another by specialized contacts. The basal layer of epidermal cells also adheres to an underlying, noncellular layer (the basement membrane). The dermis consists largely of extracellular elements that interact with each other and with the surfaces of scattered cells (primarily fibroblasts). The cells contain receptors that interact with extracellular materials and transmit signals to the cell interior.
540. 7.1: The Extracellular Space
541. The Extracellular Matrix
542. Figure 7.2: The glycocalyx. (a) The basal surface of an ectodermal cell of an early chick embryo. Two distinct structures closely applied to the outer cell surface can be distinguished: an inner glycocalyx (GC) and an outer basement membrane (BM). (b) Electron micrograph of the apical surface of an epithelial cell from the lining of the intestine, showing the extensive glycocalyx, which has been stained with the iron-containing protein ferritin.
543. Figure 7.3: The extracellular matrix (ECM) of cartilage cells. (a) Scanning electron micrograph of a portion of a colony of cartilage cells (chondrocytes) showing the extracellular materials secreted by the cells. (b) The ECM of a single chondrocyte has been made visible by adding a suspension of red blood cells (RBCs). The thickness of the ECM is evident by the clear space (arrowhead) that is not penetrated by the RBCs. The bar represents 10 μm.
544. Figure 7.4: The basement membrane (basal lamina). (a) Scanning electron micrograph of human skin. The epidermis has pulled away from part of the basement membrane, which is visible beneath the epidermal cells. (b) An unusually thick basement membrane is formed between the blood vessels of the glomerulus and the proximal end of the renal tubules of the kidney. This extracellular layer plays an important role in filtering the fluid that is forced out of the capillaries into the renal tubules during urine formation. The black dots within the glomerular basement membrane (GBM) are gold particles attached to antibodies that are bound to type IV collagen molecules in the basement membrane (CL, capillary lumen; P, podocyte of renal tubule). The bar represents 0.5 μm.
545. Collagen
546. Figure 7.5: An overview of the macromolecular organization of the extracellular matrix. The proteins and polysaccharides shown in this illustration will be discussed in the following sections. The proteins depicted (fibronectin, collagen, and laminin) contain binding sites for one another, as well as binding sites for receptors (integrins) that are located at the cell surface. The proteoglycans are huge protein–polysaccharide complexes that occupy much of the volume of the extracellular space.
547. Figure 7.6: The structure of collagen I. This figure depicts several levels of organization of a fibrillar collagen. (a) The collagen molecule (or monomer) is a triple helix composed of three helical α chains. Some types of collagen contain three identical α chains and thus are homotrimers, while others are heterotrimers containing two or three different chains. Each collagen I molecule is 295 nm in length. (b) Collagen I molecules become aligned in rows in which the molecules in one row are staggered relative to those in the neighboring row. A bundle of collagen I molecules, such as that shown here, forms a collagen fibril. The staggered arrangement of the molecules produces bands (horizontal black lines in the illustration) across the fibril that repeat with a periodicity of 67 nm (which is equal to the length of the gap between molecules plus the overlap). (c) An electron micrograph of human collagen fibrils observed after metal shadowing (see Figure 7.15). The banding pattern of the fibrils is evident. (d) An atomic force micrograph showing the surface of a collagen fibril that suggests that its subcomponents are twisted spirally around the fibril axis like a rope. The banding pattern remains evident. The arrows point to places where the fibril is slightly unwound, as would occur if you were to twist a rope in the opposite direction it is wound.
548. Figure 7.7: The corneal stroma consists largely of layers of collagen fibrils of uniform diameter and spacing. The molecules of alternate layers are arranged at right angles to one another, resembling the structure of plywood.
549. Proteoglycans
550. Figure 7.8: The type IV collagen network of the basement membrane. Electron micrograph of a basement membrane from human amniotic tissue that had been extracted with a series of salt solutions to remove noncollagenous materials. The treatment leaves behind an extensive, branching, polygonal network of strands that form an irregular lattice. Evidence indicates that this lattice consists of type IV collagen molecules covalently linked to one another in a complex three-dimensional array. A model of the basement membrane scaffold is shown in Figure 7.12.
551. Figure 7.9: The structure of a cartilage-type proteoglycan complex. (a) Schematic representation of a single proteoglycan consisting of a core protein to which a large number of glycosaminoglycan chains (GAGs, shown in red) are attached. A proteoglycan from cartilage matrix (e.g., aggrecan) may contain about 30 keratan sulfate and 100 chondroitin sulfate chains. Proteoglycans found in basement membranes (e.g., perlecan and agrin) have only a few GAG chains attached to the core protein. (b) The structures of the repeating disaccharides that make up each of the GAGs shown in this figure. All GAGs bear large numbers of negative charges (indicated by the blue shading). (c) In the cartilage matrix, individual proteoglycans are linked to a nonsulfated GAG, called hyaluronic acid (or hyaluronan), to form a giant complex with a molecular mass of about 3 million daltons. The box indicates one of the proteoglycans of the type shown in part a. (d) Electron micrograph of a proteoglycan complex, comparable to that illustrated in part c, that was isolated from cartilage matrix.
552. Fibronectin
553. Figure 7.10: Structure of fibronectin and its importance during embryonic development. (a) A human fibronectin molecule consists of two similar, but nonidentical, polypeptides joined by a pair of disulfide bonds located near the C-termini. Each polypeptide is composed of a linear series of distinct modules, which are organized into several larger functional units, illustrated by the colored cylinders in this figure. Each of these functional units contains one or more binding sites for either a specific component of the ECM or for the surface of cells. Some of these binding activities are indicated by the labels. The cell-binding site of the polypeptide that contains the sequence arg-gly-asp, or RGD, is indicated. As discussed later in the chapter, this sequence binds specifically to a particular class of integral plasma membrane proteins (integrins) that are involved in cell attachment and signal transduction. The inset shows two of the 30 or so repeating Fn modules that comprise the polypeptide; the RGD sequence forms a loop of the polypeptide, protruding from one module. (b) A section through a young chick embryo that has been treated with fluorescent antibodies against fibronectin. Fibronectin is present as fibrils in the basement membranes (dark red sites) that lie beneath the embryonic epithelia and provide a substratum over which cells migrate. (c) In this micrograph, neural crest cells are migrating out from a portion of the developing chick nervous system (beyond the edge of the photo) onto a glass culture dish that contains strips of fibronectin-coated surface alternating with strips of bare glass. The boundary of a fibronectin-coated region is indicated by the white lines. It is evident that the cells remain exclusively on the fibronectin-coated surface. Cells that reach the glass substratum (arrowheads) tend to round up and lose their migratory capabilities. The arrow indicates the direction of migration. (d, e) The role of fibronectin in the formation of the embryonic salivary gland. The micrograph in d shows a mouse embryonic salivary gland that has been grown for 10 hours in culture. The gland is seen to be divided into separate buds by a series of clefts (open triangles). The gland shown in e has been grown for the same period of time in the presence of an anti-fibronectin antibody, which has completely inhibited cleft formation. Scale bar equals 100 μm.
554. Laminin
555. Figure 7.11: The role of cell migration during embryonic development. (a) A summary of some of the cellular traffic occurring during mammalian development. The most extensive movements are conducted by the neural crest cells (shown in blue), which migrate out of the neural plate in the dorsal midline of the embryo and give rise to all the pigment cells of the skin (P), the sympathetic ganglia (SpG), adrenal medulla (AdM), and the cartilage of the embryonic skull (Mx, Md for maxillary and mandibular arches). The primordial germ cells (PGC) migrate from the yolk sac to the site of gonad (G) formation within the embryo. The progenitors of lymphoid cells are transported to the liver (L), bone marrow (Bm), thymus (Thy), lymph nodes (LN), and spleen (Sp). (Note: The “pathways” shown here connect the original sites of cells with their destinations; they do not accurately depict the actual routes of the cells.) (b) Micrograph of a section of a portion of the hind gut of a 10-day mouse embryo. The primordial germ cells (green) are seen to be migrating along the dorsal mesentery on their way to the developing gonad. The tissue has been stained with antibodies against the protein laminin (red), which is seen to be concentrated in the surface over which the cells are migrating.
556. Figure 7.12: A model of the basement membrane scaffold. Basement membranes contain two network-forming molecules, collagen IV (pink), which was illustrated in Figure 7.8, and laminin (green), which is indicated by the thickened cross-shaped molecules. The collagen and laminin networks are connected by entactin molecules (purple).
557. Dynamic Properties of the ECM
558. REVIEW
559. 7.2: Interactions of Cells with Extracellular Materials
560. Integrins
561. Table 7.1: Classification of Integrin Receptors Based on Recognition of RGD Sequences
562. Figure 7.13: Integrin conformations. (a) Ribbon drawing of a complete integrin in the bent, inactive conformation, and a corresponding electron micrograph of the extracellular portion of a similar molecule. (b) Ribbon drawing and electron micrograph of the same integrin in the extended, active (i.e., ligand-binding) conformation. Conformations that are intermediate between those shown here may also exist and exhibit reduced ligand-binding.
563. Figure 7.14: A model of integrin activation. Schematic representation of a heterodimeric integrin molecule in the bent, inactive conformation (left) and the upright, active conformation (right). The switch in conformation is triggered by the binding of a protein, in this case talin, to the small cytoplasmic domain of the β subunit. Binding of talin induces a separation of the two subunits and conversion to the active conformation. Activated integrins typically become clustered as the result of interactions of their cytoplasmic domains with the underlying cytoskeleton as indicated in Figure 7.17c. The domain structure of each subunit seen in the ribbon drawings of Figure 7.13 is shown here using globular-shaped segments. The extracellular ligand, in this case a collagen fiber, binds to both of the subunits in the head region of the activated integrin dimer.
564. Figure 7.15: The role of integrins in platelet aggregation. (a) Blood clots form when platelets adhere to one another through fibrinogen bridges that bind to the platelet integrins. (b) The presence of synthetic RGD peptides can inhibit blood clot formation by competing with fibrinogen molecules for the RGD-binding sites on platelet αIIbβ3 integrins. Nonpeptide RGD analogues and anti-integrin antibodies can act in a similar way to prevent clot formation in high-risk patients.
565. Focal Adhesions and Hemidesmosomes: Anchoring Cells to Their Substratum
566. Figure 7.16: Steps in the process of cell spreading. Scanning electron micrographs showing the morphology of mouse fibroblasts at successive times during attachment and spreading on glass coverslips. Cells were fixed after (a) 30 minutes, (b) 60 minutes, (c) 2 hours, and (d) 24 hours of attachment.
567. Figure 7.17: Focal adhesions are sites where cells adhere to their substratum and transmit signals in both directions across the plasma membrane. (a) This cultured cell has been stained with fluorescent antibodies to reveal the locations of actin filaments (gray-green) and integrins (red). The integrins are localized in small patches that correspond to the sites of focal adhesions. (b) The cytoplasmic surface of a focal adhesion of a cultured amphibian cell is shown here after the inner surface of the membrane was processed for quick-freeze, deep-etch analysis. Bundles of actin filaments are seen to associate with the inner surface of the membrane in the region of a focal adhesion. (c) Schematic drawing of a focal adhesion showing the interactions of integrin molecules with other proteins on both sides of the lipid bilayer. The binding of extracellular ligands, such as collagen and fibronectin, is thought to induce conformational changes in the cytoplasmic domains of the integrins that cause the integrins to become linked to actin filaments of the cytoskeleton. Linkages with the cytoskeleton lead, in turn, to the clustering of integrins at the cell surface. Linkages with the cytoskeleton are mediated by various actin-binding proteins, such as talin and α-actinin, that bind to the β subunit of the integrin. During the process of focal adhesion formation, talin undergoes a conformational change that exposes binding sites on talin’s rod domain. The binding of vinculin to these exposed sites promotes the assembly of additional actin filaments. The cytoplasmic domains of integrins are also associated with protein kinases, such as FAK (focal adhesion kinase) and Src. The attachment of the integrin to an extracellular ligand can activate these protein kinases and start a chain reaction that transmits signals throughout the cell. The association of myosin molecules with the actin filaments can generate traction forces that are transmitted to sites of cell–substrate attachment.
568. Figure 7.18: Experimental demonstration of forces exerted by focal adhesions. (a) In this approach, fibroblasts are plated on a deformable surface that contains a visible, uniform grid pattern. Traction forces generated by focal adhesions can be monitored by observing deformations (arrowheads) in the grid pattern of the substratum to which the cells are adhering. The generation of force can be correlated with the locations of fluorescently labeled focal adhesions (see Figure 9.73). (b) Scanning-electron micrograph of a smooth muscle cell sitting atop a bed of flexible (elastomeric) micro-posts whose tips had been coated with fibronectin. The attached cell is seen deflecting the position of multiple posts. The degree of movement of a particular post reflects the magnitude of the traction forces exerted by the cell on that post.
569. Figure 7.19: Hemidesmosomes are differentiated sites at the basal surfaces of epithelial cells where the cells are attached to the underlying basement membrane. (a) Electron micrograph of several hemidesmosomes showing the dense plaque on the inner surface of the plasma membrane and the intermediate filaments projecting into the cytoplasm. (b) Schematic diagram showing the major components of a hemidesmosome connecting the epidermis to the underlying dermis. The α6β4 integrin molecules of the epidermal cells are linked to cytoplasmic intermediate filaments by a protein called plectin that is present in the dark-staining plaque and to the basement membrane by anchoring filaments of a particular type of laminin. A second transmembrane protein (BP180) is also present in hemidesmosomes. The collagen fibers are part of the underlying dermis.
570. REVIEW
571. 7.3: Interactions of Cells with Other Cells
572. Selectins
573. Figure 7.20: Experimental demonstration of cell–cell recognition. When cells from different parts of an embryo are dissociated and then intermixed, the cells initially aggregate and then sort out by associating with other cells of the same type. The results of two such classic experiments are shown here. (a) In this experiment, two regions of an early amphibian embryo (the ectoderm and mesoderm) were dissociated into single cells and combined. At first the cells form a mixed aggregate, but eventually they sort out. The ectodermal cells (shown in red) move to the outer surface of the aggregate, which is where they would be located in the embryo, and the mesodermal cells (shown in purple) move to the interior, the position they would occupy in the embryo. Both types of cells then differentiate into the types of structures they would normally give rise to. (b) Light micrograph showing the results of an experiment in which precartilage cells from a chick limb are mixed with chick heart ventricle cells. The two types of cells have sorted themselves out of the mixed aggregate with the heart cells forming a layer on the outside of the precartilage cells. It is proposed that the precartilage cells collect in the center of the aggregate because the cells adhere to one another more strongly than do the cells from the heart. (This and other models are discussed in Nat. Cell Biol. 10:375, 2008.)
574. Figure 7.21: Selectins. Schematic drawing showing the three types of known selectins (a). All of them recognize and bind to a similar carbohydrate ligand at the ends of oligosaccharide chains on glycoproteins, such as the one shown in (b). (c) The detailed structure of the carbohydrate ligand. The terminal fucose and sialic acid moieties are particularly important in selectin recognition, and the N-acetylglucosamine moiety is often sulfated.
575. Figure 7.22: L1 is a cell-adhesion molecule of the immunoglobulin (Ig) superfamily. A model of cell–cell adhesion resulting from the specific interactions of the immunoglobulin (Ig) domains of two L1 molecules projecting from the surfaces of neighboring cells. Each L1 molecule contains a small cytoplasmic domain, a transmembrane segment, several segments that resemble one type of module found in fibronectin, and six Ig domains situated at the N-terminal portion of the molecule. The inset shows the structure of the two N-terminal Ig domains of VCAM, an IgSF molecule on the surface of endothelial cells. The Ig domains of VCAM and L1 have a similar three-dimensional structure consisting of two β sheets packed face-to-face.
576. The Immunoglobulin Superfamily
577. Cadherins
578. Figure 7.23: Cadherins and cell adhesion. Schematic representation of two cells adhering to one another as the result of interactions between similar types of cadherins projecting from the plasma membrane of each cell. Calcium ions (shown as small yellow spheres) are situated between the successive domains of the cadherin molecule where they play a critical role in maintaining the rigidity of the extracellular portion of the protein. This illustration shows several alternate models by which cadherins from opposing cells might interact. Different types of studies have suggested different degrees of overlap (interdigitation) between the extracellular domains of molecules from opposing cells. For consistency, subsequent figures will depict cadherins with a single domain overlap, which is likely the predominant configuration.
579. Figure 7.24: Cadherins and morphogenesis. (a) During gastrulation, cells in the upper layer of the embryo (the epiblast) move toward a groove in the center of the embryo, sink into the groove, and then migrate laterally as mesenchymal cells in the space beneath the epiblast. This epithelial-mesenchymal transition is marked by a loss of expression of E-cadherin that is characteristic of epithelial cells. Cells expressing E-cadherin are depicted in orange. (b) Scanning electron micrograph of a chick embryo during gastrulation that has been fractured to reveal the cells undergoing the epithelial-mesenchymal transition (arrow) depicted in part a. (c) This sequence of drawings depicts the development of the neural tube, which is an epithelial layer that forms by separation from the overlying layer of dorsal ectoderm. In the top drawing, the epithelial cells are expressing E-cadherin. In the lower drawings, the cells of the neural tube stop expressing E-cadherin (orange) and instead express N-cadherin (blue).
580. THE HUMAN PERSPECTIVE: The Role of Cell Adhesion in Inflammation and Metastasis
581. Figure 1: Steps in the movement of neutrophils from the bloodstream during inflammation. The steps are described in the text
582. Figure 2: Steps leading to the metastatic spread of an epithelial cancer (a carcinoma). (a) A fraction of the cells of the primary tumor lose their adhesiveness to other tumor cells and gain the capability to penetrate the basement membrane (BM) barrier that underlies the epithelial tissue. These cells, which have assumed a mesenchymal-like appearance, migrate through the surrounding stromal tissue and cross the BM of a blood or lymph vessel, thereby entering the general circulation. The cells are carried to other tissues, where they migrate back across the BM of the vessel and enter a tissue in which they possess the potential to form secondary tumors. Only a very small percentage of tumor cells that are released from a primary tumor manage to overcome these numerous hurdles, but those that do pose a threat to the life of the host. (b) These circulating tumor cells (CTCs) have been isolated from a blood sample of a patient with prostate cancer. Even though the blood of a cancer patient may contain less than one cancer cell for every billion normal cells, these rare cancer cells can be selectively trapped on a chip that has been coated with antibody molecules directed against a cell-surface protein (in this case, EpCAM) that is present on the cancer cells and absent from normal blood cells.
583. Adherens Junctions and Desmosomes: Anchoring Cells to Other Cells
584. Figure 7.25: An intercellular junctional complex. (a) Schematic diagram showing the junctional complex on the lateral surfaces of a simple columnar epithelial cell. The complex consists of a tight junction (zonula occludens), an adherens junction (zonula adherens), and a desmosome (macula adherens). Other desmosomes and gap junctions are located deeper along the lateral surfaces of the cells. Adherens junctions and tight junctions encircle the cell, whereas desmosomes and gap junctions are restricted to a particular site between adjacent cells. Hemidesmosomes are shown at the basal cell surface. (b) Electron micrograph of a junctional complex between two rat airway epithelial cells (TJ, tight junction; AJ, adherens junction; D, desmosome).
585. Figure 7.26: Schematic model of the molecular architecture of an adherens junction. The cytoplasmic domain of each cadherin molecule is connected to the actin filaments of the cytoskeleton by linking proteins, including β-catenin, α-catenin, and various actin-binding proteins. One of these actin-binding proteins is formin, which participates in the polymerization of the actin filaments. Actin filament assembly at the junction may be regulated by α-catenin. β-catenin is also a key element in the Wnt signaling pathway, which transmits signals from the cell surface to the cell nucleus. Disassembly of adherens junctions may release β-catenin to participate in this pathway, leading to the activation of gene expression. Another member of the catenin family, p120 catenin, binds to a site near the membrane on the cytoplasmic domain of the cadherin. p120 catenin is thought to prevent internalization of cadherins from the membrane, promote lateral interactions between cadherins, and serve as a component of intracellular signaling pathways. Numerous other proteins found in these junctions are not indicated.
586. Figure 7.27: The structure of a desmosome. (a) Electron micrograph of a desmosome from newt epidermis. (b) Schematic model of the molecular architecture of a desmosome.
587. Figure 7.28: An overview of the types of interactions involving the cell surface. Four types of cell–cell adhesive interactions are shown, as well as two types of interactions between cells and extracellular substrata. Keep in mind that the various interactions depicted here do not occur in connection with a single cell type but are shown in this manner for purposes of illustration. For example, interactions between selectins and lectins occur primarily between circulating leukocytes and the walls of blood vessels.
588. The Role of Cell-Adhesion Receptors in Transmembrane Signaling
589. REVIEW
590. 7.4: Tight Junctions: Sealing The Extracellular Space
591. Figure 7.29: The role of extracellular proteins in maintaining the differentiated state of cells. (a) These mammary gland epithelial cells were removed from a mouse and cultured in the absence of an extracellular matrix. Unlike normal differentiated mammary cells, these cells are flattened and are not engaged in the synthesis of milk proteins. (b) When extracellular matrix molecules are added back to the culture, the cells regain their differentiated appearance and synthesize milk proteins.
592. Figure 7.30: Tight junctions. (a) Electron micrograph of a section through the apical region of adjoining epithelial cells showing where the plasma membranes of the two cells come together at intermittent points within the tight junction. Inset shows the tight junction structure at higher magnification. Tight junctions block the diffusion of solutes through the paracellular pathway between cells. (b) A model of a tight junction showing the intermittent points of contact between integral proteins from two apposing membranes. Each of these contact sites extends as a paired row of proteins within the membranes, forming a barrier that blocks solutes from penetrating the space between the cells. (c) Freeze-fracture replica showing the E face of the plasma membrane of one of the cells in a region of a tight junction. The grooves in the E face are left behind after the integral membrane proteins are pulled from this half of the membrane. (d) Scanning electron micrograph of the apical surface of an epithelium showing the encircling nature of the tight junctions.
593. Figure 7.31: The molecular composition of tight junction strands. Electron micrograph of a freeze-fracture replica of cells that had been joined to one another by tight junctions. The fracture faces were incubated with two types of gold-labeled antibodies. The smaller gold particles (arrowheads) reveal the presence of claudin molecules, whereas the larger gold particles (arrows) indicate the presence of occludin. These experiments demonstrate that both proteins are present in the same tight junction strands. Bar equals 0.15 μm. The inset shows a possible arrangement of the two integral membrane proteins as they make contact in the intercellular space. Both the claudins (red) and occludin (brown) span the membrane four times.
594. REVIEW
595. 7.5: Gap Junctions and Plasmodesmata: Mediating Intercellular Communication
596. Figure 7.32: Gap junctions. (a) Electron micrograph of a section through a gap junction perpendicular to the plane of the two adjacent membranes. The “pipelines” between the two cells are seen as electron-dense beads on the apposed plasma membranes. (b) Schematic model of a gap junction showing the arrangement of six connexin subunits to form a connexon, which contains half of the channel that connects the cytoplasm of the two adjoining cells. Each connexin subunit is an integral protein with four transmembrane domains. (c) High-resolution images derived from atomic force microscopy of the extracellular surface of a single connexon in the open (left) and closed (right) conformations. Closure of the connexon was induced by exposure to elevated Ca2+ ion concentration. (d) Freeze-fracture replica of a gap junction plaque showing the large numbers of connexons and their high concentration. (The crystal structure of a gap junction can be found in Nature 458: 597, 2009.)
597. Figure 7.33: Results of an experiment demonstrating the passage of low-molecular-weight solutes through gap junctions. Micrograph showing the passage of fluorescein from one cell into which it was injected (X) to the surrounding cells.
598. Figure 7.34: Tunneling nanotubes. Scanning electron micrograph showing two cultured neuroendocrine cells connected to one another by a thin tubular process capable of carrying materials between the cytoplasm of the neighboring cells. These processes, which are only about 100 nm in diameter, are supported by an internal actin “skeleton.” The inset shows a number of fluorescently labeled vesicles caught in the act of movement between two cells.
599. Figure 7.35: Plasmodesmata. (a) Electron micrograph of a section through a plasmodesma of a fern gametophyte. The desmotubule is seen to consist of membrane that is continuous with the endoplasmic reticulum (ER) of the cytoplasm on both sides of the plasma membrane. (b) Schematic drawing of a plasmodesma. The black arrows indicate the pathways taken by molecules as they pass through the annulus from cell to cell. (c) An example of the movement of a protein from one cell to another within a plant root. The smaller inset shows the localization of the fluorescently labeled messenger RNA molecules (green) that encode a protein called Shr. The mRNA is localized within the cells of the stele (Ste), which is thus the tissue in which this protein is synthesized. The larger photo shows the localization of the fluorescently labeled Shr protein (also green), which is present both within the stele cells where it is synthesized and the adjoining endodermal cells (End) into which it has passed by way of the connecting plasmodesmata. The transported protein is localized within the nuclei of the endodermal cells where it acts as a transcription factor. Bars: 50 μm and 25 μm (inset).
600. Plasmodesmata
601. REVIEW
602. 7.6: Cell Walls
603. Figure 7.36: The plant cell wall. (a) Electron micrograph of a plant cell surrounded by its cell wall. The middle lamella is a pectin-containing layer situated between adjacent cell walls. (b) Electron micrograph showing the cellulose microfibrils and hemicellulose cross-links of an onion cell wall after extraction of the nonfibrous pectin polymers. (c) Schematic diagram of a model of a generalized plant cell wall.
604. FIGURE IN FOCUS
605. Figure 7.37: Synthesis of plant cell wall macromolecules. (a) Freeze-fracture replica of the plasma membrane of an algal cell. The rosettes are thought to represent the cellulose-synthesizing enzyme (cellulose synthase) situated within the plasma membrane. (b) A model of cellulose fibril deposition. Each rosette is thought to form a single microfibril that associates laterally with the microfibrils from other rosettes to form a larger fiber. The entire array of rosettes might move laterally within the membrane as it is pushed by the elongating cellulose molecules. Studies suggest that the direction of movement of the membrane rosettes is determined by oriented microtubules present in the cortical cytoplasm beneath the plasma membrane (discussed in Chapter 9). (c) Electron micrograph of a Golgi complex of a peripheral root cap cell stained with antibodies against a polymer of galacturonic acid, one of the major components of pectin. This material, like hemicellulose, is assembled in the Golgi complex. The antibodies have been linked to gold particles to make them visible as dark granules. The bar represents 0.25 μm.
606. REVIEW
607. Synopsis
608. Analytic Questions
609. 8: Cytoplasmic Membrane Systems: Structure, Function, and Membrane Trafficking
610. Colorized scanning electron micrograph of a human neutrophil, a type of white blood cell, ingesting a number of bacteria by the process of phagocytosis. Neutrophils are essential components of our innate immune response against pathogens.
611. Figure 8.1: Membrane-bound compartments of the cytoplasm. The cytoplasm of this root cap cell of a maize plant contains an array of membrane-bound organelles whose structure and function will be examined in this chapter. As is evident in this micrograph, the combined surface area of the cytoplasmic membranes is many times greater than that of the surrounding plasma membrane.
612. 8.1: An Overview of the Endomembrane System
613. Figure 8.2: An overview of the biosynthetic/secretory and endocytic pathways that unite endomembranes into a dynamic, interconnected network. (a) Schematic diagram illustrating the process of vesicle transport by which materials are transported from a donor compartment to a recipient compartment. Vesicles form by membrane budding, during which specific membrane proteins (green spheres) of the donor membrane are incorporated into the vesicle membrane and specific soluble proteins (purple spheres) in the donor compartment are bound to specific receptors. When the transport vesicle subsequently fuses to another membrane, the proteins of the vesicle membrane become part of the recipient membrane, and the soluble proteins become sequestered within the lumen of the recipient compartment. (b) Materials follow the biosynthetic (or secretory) pathway from the endoplasmic reticulum, through the Golgi complex, and out to various locations including lysosomes, endosomes, secretory vesicles, secretory granules, vacuoles, and the plasma membrane. Materials follow the endocytic pathway from the cell surface to the interior by way of endosomes and lysosomes, where they are generally degraded by lysosomal enzymes.
614. REVIEW
615. 8.2: A Few Approaches to the Study of Endomembranes
616. Insights Gained from Autoradiography
617. Insights Gained from the Use of the Green Fluorescent Protein
618. Figure 8.3: Autoradiography reveals the sites of synthesis and subsequent transport of secretory proteins. (a) Electron micrograph of a section of a pancreatic acinar cell that had been incubated for 3 minutes in radioactive amino acids and then immediately fixed and prepared for autoradiography. The black silver grains that appear in the emulsion following development are localized over the endoplasmic reticulum. (b–d) Diagrams of a sequence of autoradiographs showing the movement of labeled secretory proteins (represented by the silver grains in red) through a pancreatic acinar cell. When the cell is pulse-labeled for 3 minutes and immediately fixed (as shown in a), radioactivity is localized in the endoplasmic reticulum (b). After a 3-minute pulse and 17-minute chase, radioactive label is concentrated in the Golgi complex and adjacent vesicles (c). After a 3-minute pulse and 117-minute chase, radioactivity is concentrated in the secretory granules and is beginning to be released into the pancreatic ducts (d).
619. Figure 8.4: The use of green fluorescent protein (GFP) reveals the movement of proteins within a living cell. (a) Fluorescence micrograph of a live cultured mammalian cell that had been infected with the VSV virus at 40 °C. This particular strain of the VSV virus contained a VSVG gene that (1) was fused to a gene encoding the fluorescent protein GFP and (2) contained a temperature-sensitive mutation that prevented the newly synthesized VSVG protein from leaving the ER when kept at 40 °C. The green fluorescence in this micrograph is restricted to the ER. (b) Fluorescence micrograph of a live infected cell that was held at 40 °C to allow the VSVG protein to accumulate in the ER and then incubated at 32 °C for 10 minutes. The fluorescent VSVG protein has moved on to the Golgi complex. (c) Schematic drawing showing the retention of the mutant VSVG protein in the ER at 40 °C and its synchronous movement to the Golgi complex within 10 minutes of incubation at the lower temperature.
620. Insights Gained from the Biochemical Analysis of Subcellular Fractions
621. Figure 8.5: Isolation of a microsomal fraction by differential centrifugation. (a) When a cell is broken by mechanical homogenization (step 1), the various membranous organelles become fragmented and form spherical membranous vesicles. Vesicles derived from different organelles can be separated by various techniques of centrifugation. In the procedure depicted here, the cell homogenate is first subjected to low-speed centrifugation to pellet the larger particles, such as nuclei and mitochondria, leaving the smaller vesicles (microsomes) in the supernatant (step 2). The microsomes can be removed from the supernatant by centrifugation at higher speeds for longer periods of time (step 3). A crude microsomal fraction of this type can be fractionated into different vesicle types in subsequent steps. (b) Electron micrograph of a smooth microsomal fraction in which the membranous vesicles lack ribosomes. (c) Electron micrograph of a rough microsomal fraction containing ribosome-studded membranes.
622. Insights Gained from the Use of Cell-Free Systems
623. Figure 8.6: Formation of coated vesicles in a cell-free system. Electron micrograph of a liposome preparation that had been incubated with the components required to promote vesicle budding within the cell. The proteins in the medium have become attached to the surface of the liposomes and have induced the formation of protein-coated buds (arrows).
624. Insights Gained from the Study of Mutant Phenotypes
625. Figure 8.7: The use of genetic mutants in the study of secretion. (a) The first leg of the biosynthetic secretory pathway in budding yeast. The steps are described below. (b) Electron micrograph of a section through a wild-type yeast cell. (c) A yeast cell bearing a mutation in the sec12 gene whose product is involved in the formation of vesicles at the ER membrane (step 1, part a). Because vesicles cannot form, expanded ER cisternae accumulate in the cell. (d) A yeast cell bearing a mutation in the sec17 gene, whose product is involved in vesicle fusion (step 2, part a). Because they cannot fuse with Golgi membranes, the vesicles (indicated by the arrowheads) accumulate in the cell. [The mutants depicted in c and d are temperature-sensitive mutants. When kept at the lower (permissive) temperature, they are capable of normal growth and division.]
626. Figure 8.8: Inhibition of gene expression with RNA interference. (a) A control Drosophila S2 cultured cell expressing GFP-labeled mannosidase II. The fluorescent enzyme becomes localized in the Golgi complex after its synthesis in the ER. (b) A cell that has been genetically engineered to express a specific siRNA, which binds to a complementary mRNA and inhibits translation of the encoded protein. In this case, the siRNA has caused the fluorescent enzyme to remain in the ER, which has fused with the Golgi membranes. This phenotype suggests that the mRNA being affected by the siRNA encodes a protein involved in an early step of the secretory pathway during which the enzyme is synthesized in the ER and traffics to the Golgi complex. Among the genes that exhibit this phenotype when targeted by siRNAs are those encoding proteins of the COPI coat, Sar1, and Sec23. The functions of these proteins are discussed later in the chapter.
627. REVIEW
628. 8.3: The Endoplasmic Reticulum
629. Figure 8.9: The rough endoplasmic reticulum (RER). (a) Schematic diagram showing the stacks of flattened cisternae that make up the rough ER. The cytosolic surface of the ER membrane contains bound ribosomes, which gives the cisternae their rough appearance. (b) Colorized transmission electron micrograph of a portion of the rough ER of a pancreatic acinar cell. The division of the rough ER into a cisternal space (which is devoid of ribosomes) and a cytosolic space is evident. (c) Scanning electron micrograph of the rough ER in a pancreatic acinar cell. (d) Visualization of the rough ER in a whole cultured cell as revealed by immunofluorescence staining for the enzyme protein disulfide isomerase (PDI), an ER resident protein.
630. The Smooth Endoplasmic Reticulum
631. Figure 8.10: The smooth ER (SER). Electron micrograph of a Leydig cell from the testis showing the extensive smooth ER where steroid hormones are synthesized.
632. Functions of the Rough Endoplasmic Reticulum
633. Synthesis of Proteins on Membrane-Bound versus Free Ribosomes
634. Figure 8.11: Polarized structure of a secretory cell. (a) Drawing of a mucus-secreting goblet cell from the rat colon. (b) Low-power electron micrograph of a mucus-secreting cell from Brunner’s gland of the mouse small intestine. Both types of cells display a distinctly polarized arrangement of organelles, reflecting their role in secreting large quantities of mucoproteins. The basal ends of the cells contain the nucleus and rough ER. Proteins synthesized in the rough ER move into the closely associated Golgi complex and from there into membrane-bound carriers in which the final secretory product is concentrated. The apical regions of the cells are filled with secretory granules containing the mucoproteins ready for release into a duct.
635. Synthesis of Secretory, Lysosomal, or Plant Vacuolar Proteins on Membrane-Bound Ribosomes
636. FIGURE IN FOCUS
637. Figure 8.12: A schematic model of the synthesis of a secretory protein (or a lysosomal enzyme) on a membrane-bound ribosome of the RER. (a) Synthesis of the polypeptide begins on a free ribosome. As the signal sequence (shown in red) emerges from the ribosome, it binds to the SRP (step 1), which stops further translation until the SRP-ribosome-nascent chain complex can make contact with the ER membrane. The SRP-ribosome complex then collides with and binds to an SRP receptor (SR) situated within the ER membrane (step 2). Attachment of this complex to the SRP receptor is followed by release of the SRP and the association of the ribosome with a translocon of the ER membrane (step 3). These latter events are accompanied by the reciprocal hydrolyis of GTP molecules (not shown) bound to both the SRP and its receptor. In the model depicted here, the signal peptide then binds to the interior of the translocon, displacing the plug from the channel and allowing the remainder of the polypeptide to translocate through the membrane cotranslationally (step 4). After the nascent polypeptide passes into the lumen of the ER, the signal peptide is cleaved by a membrane protein (the signal peptidase, not shown), and the protein undergoes folding with the aid of ER chaperones, such as BiP. Studies suggest that translocons are organized into groups of two or four units rather than singly as shown here. (b) Cross-sectional view of the translocon channel from the side based on the X-ray crystal structure of an archaebacterial translocon. The hour-glass shape of the aqueous channel and its helical plug are evident. The ring of hydrophobic side chains (green) situated at the narrowest site within the channel is also shown. (c) Representation of a ribosome-translocon complex in the act of synthesis and translocation of a nascent protein based on cryo-EM (Section 18.8). The exit channel within the ribosome is seen to be aligned with the conducting channel within the translocon. PCC, protein conducting channel; NC, nascent chain; P-tRNA, peptidyl-t-RNA; 40S and 60S, ribosomal subunits. (An animation can be found at http://iwasa.hms.harvard.edu)
638. Processing of Newly Synthesized Proteins in the Endoplasmic Reticulum
639. Synthesis of Integral Membrane Proteins on Membrane-Bound Ribosomes
640. Figure 8.13: A schematic model for the synthesis of an integral membrane protein that contains a single transmembrane segment near the N-terminus of the nascent polypeptide. The SRP and the various components of the membrane that were shown in Figure 8.12 are also involved in the synthesis of integral proteins, but they have been omitted for simplicity. The nascent polypeptide enters the translocon just as if it were a secretory protein (step 1). However, the entry of the hydrophobic transmembrane sequence into the pore blocks further translocation of the nascent polypeptide through the channel. Steps 2–3 show the synthesis of a transmembrane protein whose N-terminus is in the lumen of the ER and C-terminus is in the cytosol. In step 2, the lateral gate of the translocon has opened and expelled the transmembrane segment into the bilayer. Step 3 shows the final disposition of the protein. Steps 2a–4a show the synthesis of a transmembrane protein whose C-terminus is in the lumen and N-terminus is in the cystosol. In step 2a, the translocon has reoriented the transmembrane segment, in keeping with its reversed positively and negatively charged flanks. In step 3a, the translocon has opened laterally and expelled the transmembrane segment into the bilayer. Step 4a shows the final disposition of the protein. White-colored + and − signs indicate the proposed charge displayed by the inner lining of the translocon. The difference in charge between the phospholipids of the cytosolic and luminal leaflets of the bilayer (indicated by the yellow + and − signs) is also thought to play a role in determining membrane protein topology. The transmembrane segments are shown as helices based on studies indicating that these regions adopt a helical secondary structure within the ribosomal exit tunnel before they enter the translocon.
641. Membrane Biosynthesis in the ER
642. Figure 8.14: Maintenance of membrane asymmetry. As each protein is synthesized in the rough ER, it becomes inserted into the lipid bilayer in a predictable orientation determined by its amino acid sequence. This orientation is maintained throughout its travels in the endomembrane system, as illustrated in this figure. The carbohydrate chains, which are first added in the ER, provide a convenient way to assess membrane sidedness because they are always present on the cisternal side of the cytoplasmic membranes, which becomes the exoplasmic side of the plasma membrane following the fusion of vesicles with the plasma membrane.
643. Synthesis of Membrane Lipids
644. Glycosylation in the Rough Endoplasmic Reticulum
645. Figure 8.15: Modifying the lipid composition of membranes. (a) Histogram indicating percentage of each of three phospholipids (phosphatidylcholine, phosphatidylserine, and sphingomyelin) in three different cellular membranes (ER, Golgi complex, and plasma membrane). The percentage of each lipid changes gradually as membrane flows from the ER to the Golgi to the plasma membrane. (b) Schematic diagram showing three distinct mechanisms that might explain how the phospholipid composition of one membrane in the endomembrane system can be different from another membrane in the system, even though the membranous compartments are spatially and temporally continuous. (1) The head groups of phospholipids of the bilayer are modified enzymatically; (2) the membrane of a forming vesicle contains a different phospholipid composition from the membrane it buds from; (3) lipids can be removed from one membrane and inserted into another membrane by lipid-transfer proteins.
646. Figure 8.16: Steps in the synthesis of the core portion of an N-linked oligosaccharide in the rough ER. The first seven sugars (five mannose and two NAG residues) are transferred one at a time to the dolichol-PP on the cytosolic side of the ER membrane (steps 1 and 2). At this stage, the dolichol with its attached oligosaccharide is then flipped across the membrane (step 3), and the remaining sugars (four mannose and three glucose residues) are attached on the luminal side of the membrane. These latter sugars are attached one at a time on the cytosolic side of the membrane to the end of a dolichol phosphate molecule (as in steps 4 and 7), which then flips across the membrane (steps 5 and 8) and donates its sugar to the growing end of the oligosaccharide chain (steps 6 and 9). Once the oligosaccharide is completely assembled, it is transferred enzymatically to an asparagine residue (within the sequence N-X-S/T) of the nascent polypeptide (step 10). The dolichol-PP is flipped back across the membrane (step 11) and is ready to begin accepting sugars again (steps 12 and 13).
647. Figure 8.17: Quality control: ensuring that misfolded proteins do not proceed forward. Based on this proposed mechanism, misfolded proteins are recognized by a glucosyltransferase (UGGT) which adds a glucose to the end of the oligosaccharide chains. Glycoproteins containing monoglucosylated oligosaccharides are recognized by the membrane-bound chaperone calnexin and given an opportunity to achieve their correctly folded (native) state. If that does not occur after repeated attempts, the protein is dislocated to the cytosol and destroyed. The steps are described in the text. A soluble chaperone (calreticulin) participates in this same quality-control pathway.
648. Mechanisms that Ensure the Destruction of Misfolded Proteins
649. Figure 8.18: A model of the mammalian unfolded protein response (UPR). The ER contains transmembrane proteins that function as sensors of stressful events that occur within the ER lumen. Under normal conditions, these sensors are present in an inactive state as the result of their association with chaperones, particularly BiP (step 1). If the number of unfolded or misfolded proteins should increase to a high level, the chaperones are recruited to aid in protein folding, which leaves the sensors in their unbound, activated state and capable of initiating a UPR. At least three distinct UPR pathways have been identified in mammalian cells, each activated by a different protein sensor. Two of these pathways are depicted in this illustration. In one of these pathways, the release of the inhibitory BiP protein leads to the dimerization of a sensor (called PERK) (step 2). In its dimeric state, PERK becomes an activated protein kinase that phosphorylates a protein (eIF2α) that is required for the initiation of protein synthesis (step 3). This translation factor is inactive in the phosphorylated state, which stops the cell from synthesizing additional proteins in the ER (step 4), giving the cell more time to process those proteins already present in the ER lumen. In the second pathway depicted here, release of the inhibitory BiP protein allows the sensor (called ATF6) to move on to the Golgi complex where the cytosolic domain of the protein is cleaved away from its transmembrane domain (step 2a). The cytosolic portion of the sensor diffuses through the cytosol (step 3a) and into the nucleus (step 4a), where it stimulates the expression of genes whose encoded proteins can alleviate the stress in the ER (step 5a). These include chaperones, coat proteins that form on transport vesicles, and proteins of the quality-control machinery. (Discussion of the third protein sensor (IRE1) can be found in Science 334:1081, 2011.)
650. From the ER to the Golgi Complex: The First Step in Vesicular Transport
651. Figure 8.19: Visualizing membrane traffic with the use of a fluorescent tag This series of photographs shows a small portion of a living mammalian cell that has been infected with the vesicular stomatitis virus (VSV) containing a VSVG-GFP chimeric gene (page 274). Once it is synthesized in the RER, the fusion protein emits a green fluorescence, which can be followed as the protein moves through the cell. In the series of photographs shown here, two vesicular-tubular carriers (VTCs) (arrows) containing the fluorescent protein have budded from the ER and are moving toward the Golgi complex (GC). The series of events depicted here took place over a period of 13 seconds. Bar represents 6 μm.
652. REVIEW
653. 8.4: The Golgi Complex
654. Figure 8.20: The Golgi complex. (opposite) (a) Schematic model of a portion of a Golgi complex from an epithelial cell of the male rat reproductive tract. The elements of the cis and trans compartments are often discontinuous and appear as tubular networks. (b) Electron micrograph of a portion of a tobacco root cap cell showing the cis to trans polarity of the Golgi stack. (c) Electron tomographic image of a slice from a mouse pancreatic beta cell that synthesizes and secretes the protein insulin. The individual Golgi stacks are seen to be interconnected to form a continuous ribbon. The trans face (or TGN) of each Golgi stack has been colored red and the cis face has been colored light blue. Cellular tomography is discussed in Section 18.2. (d) Fluorescence micrograph of a cultured mammalian cell. The position of the Golgi complex is revealed by the red fluorescence, which marks the localization of antibodies to a COPI coat protein. (e) Electron micrograph of a single isolated Golgi cisterna showing two distinct domains, a concave central domain and an irregular peripheral domain. The peripheral domain consists of a tubular network from which protein-coated buds are being pinched off.
655. Figure 8.21: Regional differences in membrane composition across the Golgi stack. (a) Reduced osmium tetroxide preferentially impregnates the cis cisternae of the Golgi complex. (b) The enzyme mannosidase II, which is involved in trimming the mannose residues from the core oligosaccharide as described in the text, is preferentially localized in the medial cisternae. (c) The enzyme nucleoside diphosphatase, which splits dinucleotides (e.g., UDP) after they have donated their sugar, is preferentially localized in the trans cisternae.
656. Glycosylation in the Golgi Complex
657. The Movement of Materials through the Golgi Complex
658. Figure 8.22: Steps in the glycosylation of a typical mammalian N-linked oligosaccharide in the Golgi complex. Following the removal of the three glucose residues, various mannose residues are subsequently removed, while a variety of sugars (N-acetylglucosamine, galactose, fucose, and sialic acid) are added to the oligosaccharide by specific glycosyltransferases. These enzymes are integral membrane proteins whose active sites face the lumen of the Golgi cisternae. This is only one of numerous glycosylation pathways.
659. Figure 8.23: The dynamics of transport through the Golgi complex. (a) In the vesicular transport model, cargo (black dots) is carried in an anterograde direction by transport vesicles, while the cisternae themselves remain as stable elements. (b) In the cisternal maturation model, the cisternae progress gradually from a cis to a trans position and then disperse at the TGN. Transport vesicles carry resident Golgi enzymes (indicated by the colored vesicles) in a retrograde direction. The red lens-shaped objects represent large cargo materials, such as procollagen complexes of fibroblasts. (c) Electron micrograph of an area of Golgi complex in a thin frozen section of a cell that had been infected with vesicular stomatitis virus (VSV). The black dots are nanosized gold particles bound by means of antibodies to VSVG protein, an anterograde cargo molecule. The cargo is restricted to the cisternae and does not appear in nearby vesicles (arrows). (d) Electron micrograph of similar nature to that of c but, in this case, the gold particles are not bound to cargo, but to mannosidase II, a resident enzyme of the medial Golgi cisternae. The enzyme appears in both a vesicle (arrow) and cisternae. The labeled vesicle is presumably carrying the enzyme in a retrograde direction, which compensates for the anterograde movement of the enzyme as the result of cisternal maturation. Bar, 0.2 μm. (A third model for intra-Golgi transport is discussed in Cell 133:951, 2008.)
660. REVIEW
661. 8.5: Types of Vesicle Transport and Their Functions
662. Figure 8.24: Coated vesicles. These electron micrographs show the membranes of these vesicles to be covered on their outer (cytosolic) surface by a distinct protein coat. The micrograph on the left (a) shows a COPII-coated vesicle, whereas the micrograph on the right (b) shows a COPI-coated vesicle.
663. COPII-Coated Vesicles: Transporting Cargo from the ER to the Golgi Complex
664. Figure 8.25: Proposed movement of materials by vesicular transport between membranous compartments of the biosynthetic/secretory pathway. (a) The three different types of coated vesicles indicated in this schematic drawing are thought to have distinct transport roles. COPII-coated vesicles mediate transport from the ER to the ERGIC and Golgi complex. COPI-coated vesicles return proteins from the ERGIC and Golgi complex to the ER. COPI-coated vesicles also transport Golgi enzymes between cisternae in a retrograde direction. Clathrin-coated vesicles mediate transport from the TGN to endosomes and lysosomes. Transport of materials along the endocytic pathway is not shown in this drawing. (b) Schematic drawing of the assembly of a COPII-coated vesicle at the ER. Assembly begins when Sar1 is recruited to the ER membrane and activated by exchange of its bound GDP with a bound GTP. These steps are shown in Figure 8.26. Cargo proteins of the ER lumen (red spheres and diamonds) bind to the luminal ends of transmembrane cargo receptors. These receptors are then concentrated within the coated vesicle through interaction of their cytosolic tails with components of the COPII coat. ER resident proteins (e.g., BiP) are generally excluded from the coated vesicles. Those that do happen to become included in a coated vesicle are returned to the ER as described later in the text.
665. Figure 8.26: Proposed roles of the COPII coat proteins in generating membrane curvature, assembling the protein coat, and capturing cargo. In step 1, Sar1-GDP molecules have been recruited to the ER membrane by a protein called a GEF (guanine-exchange factor) that catalyzes the exchange of the bound GDP with a bound GTP. In step 2, each Sar1-GTP molecule has extended a finger-like α helix along the membrane within the cytosolic leaflet. This event expands the leaflet and induces the curvature of the lipid bilayer at that site. In step 3, a dimer composed of two COPII polypeptides (Sec23 and Sec24) has been recruited by the bound Sar1-GTP. The Sec23-Sec24 heterodimer is thought to further induce the curvature of the membrane in the formation of a vesicle. Both Sar1 and Sec23-Sec24 can bring about membrane curvature when incubated with synthetic liposomes in vitro. Transmembrane cargo accumulates within the forming COPII vesicle as their cytosolic tails bind to the Sec24 polypeptide of the COPII coat. Sec24 can exist in at least four different isoforms. It is likely that different isoforms of this protein recognize and bind membrane proteins with different sorting signals, thus broadening the specificity in types of materials that can be transported by COPII vesicles. In step 4, the remaining COPII polypeptides (Sec13 and Sec31) have joined the complex to form an outer structural scaffold of the coat.
666. Figure 8.27: A molecular model of the outer Sec13-Sec31 cage of the COPII coat as it would assemble around the surface of a 40-nm “vesicle.” Each edge or leg of the lattice that makes up the cage consists of a heterotetramer (two Sec31 subunits seen as dark green and light green and two Sec13 subunits seen as orange and red). Four such legs converge to form each vertex of the lattice. Two copies of the Sar1-Sec23-Sec24 complex (shown in red, magenta, and blue, respectively) that would form the inner layer of the COPII coat are also shown in this model. It can be seen how the inner surface of the Sec23-Sec24 complex conforms to the curvature of the vesicle. Inset shows a COPII lattice, which is comprised of triangular and square, pentagonal, and/or hexagonal faces.
667. COPI-Coated Vesicles: Transporting Escaped Proteins Back to the ER
668. Retaining and Retrieving Resident ER Proteins
669. Figure 8.28: Retrieving ER proteins. Resident proteins of the ER contain amino acid sequences that lead to their retrieval from the Golgi complex if they are accidentally incorporated into a Golgi-bound transport vesicle. Soluble ER proteins bear the retrieval signal KDEL. Retrieval is accomplished as soluble ER proteins bind to KDEL receptors residing in the membranous wall of cis Golgi compartments. The KDEL receptors, in turn, bind to proteins of the COPI coat, which allows the entire complex to be recycled back to the ER.
670. Beyond the Golgi Complex: Sorting Proteins at the TGN
671. Sorting and Transport of Lysosomal Enzymes
672. Figure 8.29: Targeting lysosomal enzymes to lysosomes. (a) Lysosomal enzymes are recognized by an enzyme in the cis cisternae that transfers a phosphorylated N-acetylglucosamine from a nucleotide sugar donor to one or more mannose residues of N-linked oligosaccharides. The glucosamine moiety is then removed in a second step by a second enzyme, leaving mannose 6-phosphate residues as part of the oligosaccharide chain. (b) Schematic diagram showing the pathways followed by a lysosomal enzyme (black) from its site of synthesis in the ER to its delivery to a lysosome. The mannose residues of the lysosomal enzyme are phosphorylated in the Golgi cisternae (step 1) and then selectively incorporated into a clathrin-coated vesicle at the TGN (step 2). The mannose 6-phosphate receptors are thought to have a dual role (step 3): they interact specifically with the lysosomal enzymes on the luminal side of the vesicle, and they interact specifically with adaptors on the cytosolic surface of the vesicle (shown in Figure 8.30). The mannose 6-phosphate receptors separate from the enzymes (step 4) and are returned to the Golgi complex (step 5). The lysosomal enzymes are delivered to an endosome (step 6) and eventually to a lysosome. Mannose 6-phosphate receptors are also present in the plasma membrane, where they capture lysosomal enzymes that are secreted into the extracellular space and return the enzymes to a pathway that directs them to a lysosome (step 7).
673. Figure 8.30: The formation of clathrin-coated vesicles at the TGN. Clathrin-coated vesicles that bud from the TGN contain GGA, an adaptor protein consisting of several distinct domains. One of the GGA domains binds to the cytosolic domains of membrane proteins, including those that will ultimately reside in the boundary membrane of the lysosome and also the MPR that transports lysosomal enzymes. Other GGA domains bind to Arf1 and to the surrounding cytosolic network of clathrin molecules.
674. Sorting and Transport of Nonlysosomal Proteins
675. Targeting Vesicles to a Particular Compartment
676. Figure 8.31: Proposed steps in the targeting of transport vesicles to target membranes. (a) According to this model, Rab proteins on the vesicle and target membrane are involved in recruiting tethering proteins that mediate initial contact between the two membranes. Two types of tethering proteins are depicted: highly elongated fibrous proteins (e.g., golgins and EEA1) and multiprotein complexes (e.g., the exocyst and TRAPPI). (b) Electron tomographic image of a slice through a mammalian nerve terminus showing the network of synaptic vesicles that are present in close association with the presynaptic plasma membrane (page 169). The left insets (corresponding to the white box on the right) show the presence of filamentous connectors between a pair of synaptic vesicles (upper inset) and between one of the synaptic vesicles and the adjacent plasma membrane (lower inset). Bars: main panel, 100 nm; insets, 50 nm. (c) During the docking stage leading up to membrane fusion, a v-SNARE in the vesicle membrane interacts with the t-SNAREs in the target membrane to form a four-stranded α-helical bundle that brings the two membranes into intimate contact (see next figure). In the cases described in the text, SNAP-25, one of the t-SNAREs, is a peripheral membrane protein that is bound to the lipid bilayer by a lipid anchor rather than a transmembrane domain. SNAP-25 contributes two helices to the four-helix SNARE bundle. (d) A model of a synaptic vesicle showing the distribution of only one of its constitutent proteins, the SNARE synaptobrevin. The surface density and structures of the synaptobrevin molecules are based on calculations of the number of these proteins per vesicle and the known structure of the molecule. A complete portrait of the proteins on the surface of a synaptic vesicle is shown in the image in Figure 4.4d.
677. Exocytosis
678. Figure 8.32: A model of the interactions between v- and t-SNAREs leading to membrane fusion and exocytosis. (a) The synaptic vesicle has become docked to the plasma membrane through the formation of four-stranded bundles comprising α helices donated by syntaxin (red), synaptobrevin (blue), and SNAP-25 (green). SNAP-25 contributes two helices and lacks a transmembrane domain (yellow). (b) A speculative transition state in the fusion of the two membranes. A small water-filled cavity is shown at the center of the transmembrane helix bundle. (c) The transmembrane helices that previously resided in the two separate membranes are now present in the same bilayer, and a fusion pore has opened between the vesicle and target membrane. The neurotransmitter contents of the vesicle can now be discharged by exocytosis. (d) Scanning electron micrograph of the extracellular surface of a pair of cultured alveolar (lung) cells stimulated to discharge proteins that had been stored in secretory granules. Material is seen being expelled from the cell through smooth, circular openings that are presumed to be dilated fusion pores. The arrow shows a fusion pore that has not dilated, which is readily distinguished from holes (arrowhead) accidentally formed during specimen preparation.
679. REVIEW
680. 8.6: Lysosomes
681. Table 8.1: A Sampling of Lysosomal Enzymes
682. Figure 8.33: Lysosomes. Portion of a phagocytic Kupffer cell of the liver showing at least 10 lysosomes of highly variable size.
683. Autophagy
684. Figure 8.34: Autophagy. Electron micrograph of a mitochondrion and peroxisome enclosed in a double membrane wrapper. This autophagic vacuole (or autophagosome) would have fused with a lysosome and its contents digested.
685. Figure 8.35: A summary of the autophagic pathway. The steps are described in the text.
686. REVIEW
687. THE HUMAN PERSPECTIVE: Disorders Resulting from Defects in Lysosomal Function
688. Figure 1: Lysosomal storage disorders. Electron micrograph of a section through a portion of a neuron of a person with a lysosomal storage disease characterized by an inability to degrade GM2 gangliosides. These cytoplasmic vacuoles stain for both lysosomal enzymes and the ganglioside, indicating they are lysosomes in which undigested glycolipids have accumulated.
689. Table 1: Sphingolipid Storage Diseases
690. Figure 8.36: Plant cell vacuoles. (a) Each of the cylindrical leaf cells of the aquatic plant Elodea contains a large central vacuole surrounded by a layer of cytoplasm containing the chloroplasts that are visible in the micrograph. (b) Transmission electron micrograph of spinach leaf mesophyll cells showing the large central vacuole and thin layer of surrounding cytoplasm.
691. 8.7: Plant Cell Vacuoles
692. REVIEW
693. 8.8: The Endocytic Pathway: Moving Membrane and Materials into the Cell Interior
694. Endocytosis
695. Receptor-Mediated Endocytosis and the Role of Coated Pits
696. Figure 8.37: Receptor-mediated endocytosis. This sequence of micrographs shows the steps in the uptake of yolk lipoproteins by the hen oocyte. (a) The proteins to be taken into the cell are concentrated on the extracellular surface of an indented region of the plasma membrane, forming a coated pit. The cytosolic surface of the plasma membrane of the coated pit is covered with a layer of bristly, electron-dense material containing the protein clathrin. (b) The coated pit has sunk inward to form a coated bud. (c) The plasma membrane is about to pinch off as a vesicle containing the yolk protein on its luminal (previously extracellular) surface and clathrin on its cytosolic surface. (d) A coated vesicle that is no longer attached to the plasma membrane. The next step in the process is the release of the clathrin coat.
697. Figure 8.38: Coated pits. (a) Electron micrograph of a replica formed on the extracellular surface of a whole, freeze-dried fibroblast that had been incubated with LDL-cholesterol. Particles of LDL-cholesterol are visible as spherical structures located on the extracellular surface of the coated pit. (b) Electron micrograph of a replica formed on the cytosolic surface of a coated pit of a ruptured fibroblast. The coat is composed of a flattened network of clathrin-containing polygons associated with the inner surface of the plasma membrane. (c) Electron micrograph of a replica of the cytosolic surface of a coated bud showing the invaginated plasma membrane surrounded by a clathrin lattice that has assumed a hemispheric shape.
698. Figure 8.39: Clathrin triskelions. Electron micrograph of a metal-shadowed preparation of clathrin triskelions. Inset shows the triskelion is composed of three heavy chains. The inner portion of each heavy chain is linked to a smaller light chain.
699. Figure 8.40: Molecular organization of a coated vesicle. (a) Schematic drawing of the surface of a coated vesicle showing the arrangement of triskelions and adaptors in the outer clathrin coat. The sides of the polygons are formed by parts of the legs of overlapping triskelions. The N-terminus of each clathrin heavy chain forms a “hook” that projects toward the surface of the membrane where it engages an adaptor. Each adaptor, which consists of four different polypeptide subunits, can bind a diverse array of accessory proteins that are not shown in this illustration. Both the hooks and adaptors are situated at the vertices of the polyhedrons. (Note: Not all of the triskelions of the lattice are shown in this figure; if they were, every vertex would have a clathrin hub, hook, and associated adaptor.) (b) Schematic drawing of a cross section through the surface of a coated vesicle showing the interactions of the AP2 adaptor complexes with both the clathrin coat and membrane receptors. Recruitment of AP2 adaptors to the plasma membrane is facilitated by the presence of PI(4,5)P2 molecules in the inner (cytosolic) leaflet of the membrane as shown in Figure 8.42. Each receptor is bound to a ligand being internalized. (c) Reconstruction of a clathrin cage containing 36 triskelions showing the overlapping arrangement of several of these trimeric molecules (shown in different colors). (An animation can be found at http://iwasa.hms.harvard.edu)
700. Figure 8.41: The role of dynamin in the formation of clathrin-coated vesicles. (a) The clathrin lattice of the coated pit (step 1) undergoes rearrangement to form an invaginated vesicle connected to the overlying plasma membrane by a stalk (step 2). At this point, the dynamin subunits, which are concentrated in the region, undergo polymerization to form a ring around the stalk (step 3). Changes in the conformation of the ring, which are thought to be induced by GTP hydrolysis (step 4), lead to fission of the coated vesicle from the plasma membrane and disassembly of the dynamin ring (step 5a). If vesicle budding occurs in the presence of GTPγS, a nonhydrolyzable analogue of GTP, dynamin polymerization continues beyond formation of a simple collar, producing a narrow tubule constructed from several turns of the dynamin helix (step 5b). Structural models of dynamin action can be found in Nature 477:556, 561, 2011. (b) Electron micrograph showing a coated vesicle forming in the presence of GTPγS, which corresponds to the stage depicted in step 5b of part a.
701. The Role of Phosphoinositides in the Formation of Coated Vesicles
702. Figure 8.42: A structural model depicting the changes in protein conformation that occur upon AP2 binding to the plasma membrane. Binding of the AP2 adaptor is mediated by its interaction with PI(4,5)P2 molecules in the inner leaflet of the plasma membrane (step 1). Binding of the adaptor to the membrane induces a large conformational change in the adaptor (step 2) that facilitates its interaction with specific motifs in the cytoplasmic tails of certain membrane receptors (shown in yellow and orange) (step 3).
703. The Endocytic Pathway
704. Figure 8.43: The endocytic pathway. The movement of materials from the extracellular space to early endosomes where sorting occurs. Endocytosis of two types of receptor–ligand complexes is shown. Housekeeping receptors, such as the LDL receptor (shown in red), are typically sent back to the plasma membrane, whereas their ligands (purple spheres) are transferred to late endosomes. Signaling receptors, such as the EGF receptor (shown in green), are typically transported to late endosomes along with their ligands (yellow). Late endosomes also receive newly synthesized lysosomal enzymes (red spheres) from the TGN. These enzymes are carried by mannose 6-phosphate receptors (MPRs), which return to the TGN. The contents of late endosomes are transferred to lysosomes by a number of routes (not shown). The inset on the left shows an enlarged view of a portion of a late endosome with an intraluminal vesicle budding inward from the outer membrane. The membranes of these vesicles contain receptors to be degraded. (b) Electron micrograph showing the internal vesicles within the lumen of a late endosome. A number of lysosomes are present in the vicinity. (c) The gold particles seen in this electron micrograph are bound to EGF receptors that were internalized by endocytosis and have become localized within the membranes of the internal vesicles of this late endosome.
705. LDLs and Cholesterol Metabolism
706. Figure 8.44: LDL cholesterol. Each particle consists of esterified cholesterol molecules, surrounded by a mixed monomolecular layer of phospholipids and cholesterol, and a single molecule of the protein apolipoprotein B-100, which interacts specifically with the LDL receptor projecting from the plasma membrane.
707. Figure 8.45: A model of atherosclerotic plaque formation. According to this model, plaque formation is initiated by various types of injury to the endothelial cells that line the vessel, including damage inflicted by oxygen free radicals that chemically alter the LDL-cholesterol particles. The injured endothelium acts as an attractant for white blood cells (leukocytes) and macrophages, which migrate beneath the endothelium and begin a process of chronic inflammation. The macrophages ingest the oxidized LDL, which becomes deposited in the cytoplasm as cholesterol-rich fatty droplets. These cells are referred to as macrophage foam cells and they are often already present in the blood vessels of adolescents and young adults. Substances released by the macrophages stimulate the proliferation of smooth muscle cells, which produce a dense, fibrous connective tissue matrix (fibrous cap) that bulges into the arterial lumen. Not only do these bulging lesions restrict blood flow, they are prone to rupture, which can trigger the formation of a blood clot and ensuing heart attack.
708. Figure 8.46: Phagocytosis. (a) The process of engulfment as illustrated by a polymorphonuclear leukocyte ingesting a yeast cell (lower left). (b) The steps that occur in the phagocytic pathway (see also page 305).
709. Phagocytosis
710. REVIEW
711. 8.9: Posttranslational Uptake of Proteins by Peroxisomes, Mitochondria, and Chloroplasts
712. Uptake of Proteins into Peroxisomes
713. Uptake of Proteins into Mitochondria
714. Figure 8.47: Importing proteins into a mitochondrion. (a) Proposed steps taken by proteins imported posttranslationally into either the mitochondrial matrix or inner mitochondrial membrane. The polypeptide is targeted to a mitochondrion by a targeting sequence, which is located at the N-terminus in the matrix protein (step 1) and is located internally in most inner membrane proteins (step A). Cytosolic Hsp70 molecules unfold the polypeptides prior to their entry into the mitochondrion. The proteins are recognized by membrane receptors (red transmembrane proteins) and translocated through the OMM by way of pores in the TOM complex of the OMM (step 2 or B). Most integral proteins of the IMM are directed to the TIM22 complex of the IMM (step C), which steers them into the lipid bilayer of the IMM (step D). Mitochondrial matrix proteins are translocated through the TIM23 complex of the IMM (step 3). Once the protein enters the matrix, it is bound by a mitochondrial chaperone (step 4), which may either pull the polypeptide into the matrix or act like a Brownian ratchet to ensure that it diffuses into the matrix (these alternate chaperone mechanisms are discussed in the text). Once in the matrix, the unfolded protein assumes its native conformation (step 5a) with the help of Hsp60 chaperones (not shown). The presequence is removed enzymatically (step 5b). (b) A three-dimensional model of the mitochondrial protein-import machinery, showing the number, relative size, and topology of the various proteins involved in this activity. The TOM complex is a reddish color, the TIM23 complex is yellow-green, the TIM22 complex is green, and the cooperating chaperones are blue.
715. Uptake of Proteins into Chloroplasts
716. Figure 8.48: Importing proteins into a chloroplast. Proteins encoded by nuclear genes are synthesized in the cytosol and imported through protein-lined pores in both membranes of the outer chloroplast envelope (step 1). Proteins destined for the stroma (step 1a) contain a stroma-targeting domain at their N-terminus, whereas proteins destined for the thylakoid (step 1b) contain both a stroma-targeting domain and a thylakoid-transfer domain at their N-terminus. Stromal proteins remain in the stroma (step 2) following translocation through the outer envelope and removal of their single targeting sequence. The presence of the thylakoid transfer domain causes thylakoid proteins to be translocated either into or completely through the thylakoid membrane (step 3). A number of the proteins of the thylakoid membrane are encoded by chloroplast genes and synthesized by chloroplast ribosomes that are bound to the outer surface of the thylakoid membrane (step 4).
717. REVIEW
718. EXPERIMENTAL PATHWAYS: Receptor-Mediated Endocytosis
719. Figure 1: (a) A handmade model of an “empty basket” that would form the surface lattice of a coated vesicle. (b) A high-power electron micrograph of an empty proteinaceous basket. The numbers 5 and 6 refer to pentagonal and hexagonal elements of the lattice, respectively.
720. Figure 2: HMG CoA reductase activity in normal fibroblasts was measured following addition of the lipoprotein fraction of calf serum (open squares), unfractionated calf serum (closed circles), or nonlipoprotein fraction of calf serum (open triangles). It is evident that the lipoproteins greatly depress the activity of the enzyme, while the nonlipoproteins have little effect.
721. Figure 3: Fibroblast cells from either a control subject (closed circles) or a patient with homozygous FH (open circles) were grown in dishes containing fetal calf serum. On day 6 (which corresponds to 0 hours on the graph), the medium was replaced with fresh medium containing lipoprotein-deficient human plasma. At the indicated time, extracts were prepared, and HMG CoA reductase activity was measured. If we look at the control cells, it is apparent that at the beginning of the monitoring period the cells have very little enzyme activity because the medium had contained enough cholesterol-containing lipoproteins that the cells did not need to synthesize their own. Once the medium was changed to the lipoprotein-deficient plasma, the cells were no longer able to use cholesterol from the medium and thus increased the amount of enzyme within the cell. In contrast, the cells from the FH patients showed no response to either the presence or absence of lipoproteins in the medium.
722. Figure 4: Time course of radioactive [125I]-labeled LDL binding to cells from a normal subject (circles) and a homozygote with FH (triangles) at 37 °C. The cells were incubated in a buffer containing 5 mg/ml [125I] LDL in the presence (open circles and triangles) and absence (closed circles and triangles) of 250 mg/ml of nonradioactive LDL. It is evident that in the absence of added nonradioactive LDL, the normal cells bind significant amounts of the labeled LDL, while the cells from the FH patients do not. The binding of labeled LDL is greatly reduced in the presence of nonradioactive LDL because the nonlabeled lipoproteins compete with the labeled ones for binding sites. Thus, the binding of the lipoprotein to the cells is specific (i.e., it is not the result of some nonspecific binding event).
723. Figure 5: Electron micrograph showing the binding of LDL to a coated pit of a human fibroblast. The LDL is made visible by conjugating the particles to electron-dense, iron-containing ferritin. Bar: 100 nm.
724. Figure 6: A series of fluorescence images showing the capture of a single red-fluorescent LDL particle by a green-fluorescent clathrin-coated pit and its incorporation into a clathrin-coated vesicle, which becomes uncoated and moves into the cytoplasm. The events are described in the text. T1-T4 represent the intervals before association (T1), during association (T2), joint lateral motion of LDL and clathrin prior to uncoating (T3), and motion of LDL after uncoating (T4), respectively. The times are indicated in seconds in each frame, with 0 corresponding to the time when the LDL and clathrin become associated.
725. References
726. Synopsis
727. Analytic Questions
728. 9: The Cytoskeleton and Cell Motility
729. The cytoskeleton is constructed from three major types of filaments, which are shown imaged separately in parts a–c. The distribution of each type of filament within a cultured liver cell (hepatocyte) is revealed by a fluorescent probe that binds specifically to that type of filament. The distribution of microfilaments (a) is revealed by fluorescent phalloidin binding; the distribution of microtubules (b) by fluorescent anti-tubulin antibodies; and the distribution of intermediate filaments (c) by fluorescent anti-keratin antibodies. The image in d is a composite of the three previous images and demonstrates that the cytoplasmic organization of each filament type is distinct. Bar equals 10 μm.
730. Table 9.1: Properties of Microtubules, Intermediate Filaments, and Actin Filaments
731. 9.1: Overview of the Major Functions of the Cytoskeleton
732. Figure 9.1: An overview of the structure and functions of the cytoskeleton. Schematic drawings of (a) an epithelial cell, (b) a nerve cell, and (c) a dividing cell. The microtubules of the epithelial and nerve cells function primarily in support and organelle transport, whereas the microtubules of the dividing cell form the mitotic spindle required for chromosome segregation. Intermediate filaments provide structural support for both the epithelial cell and nerve cell. Microfilaments support the microvilli of the epithelial cell and are an integral part of the motile machinery involved in nerve cell elongation and cell division.
733. Figure 9.2: An example of the role of microtubules in transporting organelles. The peroxisomes of this cell (shown in green and indicated by arrows) are closely associated with microtubules of the cytoskeleton (shown in red). Peroxisomes appear green because they contain a peroxisomal protein fused to the green fluorescent protein (page 273). Microtubules appear red because they are stained with a fluorescently labeled antibody.
734. 9.2: The Study of the Cytoskeleton
735. The Use of Live-Cell Fluorescence Imaging
736. Figure 9.3: The capability of the cytoskeleton to change its three-dimensional organization is apparent in this mouse fibroblast that is migrating over the 90° edge of a coverslip. Bar, 30 μm.
737. Figure 9.4: Dynamic changes in length of microtubules within an epithelial cell. The cell was injected with a small volume of tubulin that had been covalently linked to the fluorescent dye rhodamine. After allowing time for the cell to incorporate the labeled tubulin into microtubules, a small portion of the edge of the living cell was examined with the fluorescence microscope. The elapsed time is shown at the lower right of each image. The microtubules have been false-colored to increase their contrast. A “pioneering” microtubule (arrowhead) can be seen to grow out to the leading edge of the cell, where it becomes bent and then continues its growth in a direction parallel to the cell’s edge. The rate of growth of bent parallel microtubules was much greater than those growing perpendicular to the cell’s edge. Bar, 10 μm
738. The Use of In Vitro and In Vivo Single-Molecule Assays
739. Figure 9.5: The localization of a protein within a cell using fluorescent antibodies. This algal cell was stained with fluorescent antibodies (yellow-green color) directed against a protein called centrin. Centrin is seen to be localized within the cell’s flagella and the rootlike structure at the base of the flagella. The red color of the cell results from autofluorescence of chlorophyll molecules of this photosynthetic alga.
740. Figure 9.6: Following the movements of individual, fluorescently labeled kinesin molecules along microtubules both in vitro and in vivo. (a) In this experiment a kinesin molecule labeled with a GFP variant called mCit (green) is seen to move processively along a microtubule whose plus end is labeled with a red fluorescent dye. The successive images, from left to right, represent separate micrographs taken at increasing time periods over the course of the experiment. The kinesin is moving at an average speed of 0.77 μm per second. (The intensity of the green-colored fluorescence diminishes over time as the result of photobleaching that occurs during observation.) Bar, 2 μm. (b) Schematic illustration of the experiment shown in part a. In the actual experiment, more than one mCit molecule is bound per kinesin molecule. (c) The image shows a portion of a living cell that has synthesized mCit-labeled kinesin molecules (yellow-green). Several of the kinesin molecules can be seen bound to the microtubules. The image seen here represents a single point in time. When observed over a time period, the kinesin molecules are seen to scurry along the microtubules. (Kinesin movement can be seen by watching movie 6 of this paper at www.biophysj.org)
741. Figure 9.7: Determining the mechanical properties of single cytoskeletal filaments. These images show a single intermediate filament before and after it has been subjected to mechanical forces by the tip of an atomic force microscope. The tip is attached at one point along the filament (arrow) and has moved to the left, pulling on the affected segment. This segment has been stretched more than three times its original length (from 280 nm to a stretched length of 500+450 nm).
742. The Use of Fluorescence Imaging Techniques to Monitor the Dynamics of the Cytoskeleton
743. Figure 9.8: The study of the cytoskeleton using FRAP. (a) A cell expressing GFP-tubulin incorporates the fluorescent tubulin into its microtubule array. When a laser is focused on a box on the array, the fluorescence is bleached (t=0 sec). Over time the fluorescence is recovered in the bleached zone. (b) The fluorescence recovery can come from different properties of microtubules. The dynamic properties of the microtubules are very likely to contribute to fluorescence recovery. Alternatively, if new fluorescent microtubules grow from the centrosome, they can grow into the bleached zone. Finally, it is possible, that a fluorescent translocating microtubule can move through the bleached zone and be seen there at the time of observation. (c) Micrographs from a FRAP experiment performed on interphase (non-dividing) cells expressing GFP-tubulin. Two side-by-side cells were bleached in the boxed region with a laser, and the cells were then imaged over time. The fluorescence signal in the bleached region recovers over the time course of the experiment.
744. REVIEW
745. 9.3: Microtubules
746. Structure and Composition
747. FIGURE IN FOCUS
748. Figure 9.9: The structure of microtubules. (a) Electron micrograph of negatively stained microtubules from brain showing the globular subunits that make up the protofilaments. The bumps at the surface of the microtubules are microtubule-associated proteins (MAPs), discussed later. (b) Electron micrograph of a cross section through a microtubule of a Juniperus root tip cell revealing the 13 subunits arranged within the wall of the tubule. The microtubules of these plant cells are most abundant in a cortical zone about 100 nm thick just beneath the plasma membrane (seen in the lower right of the micrograph). (c) A ribbon model showing the three-dimensional structure of the αβ-tubulin heterodimer. Note the complementary shapes of the subunits at their interacting surfaces. The α-tubulin subunit has a bound GTP, which is not hydrolyzed and is nonexchangeable. The β-tubulin subunit has a bound GDP, which is exchanged for a GTP prior to assembly into a polymer (page 342). The plus end of the dimer is at the top. (d) Diagram of a longitudinal section of a microtubule shown in the B-lattice, which is the structure thought to occur in the cell. The wall consists of 13 protofilaments composed of αβ-tubulin heterodimers stacked in a head-to-tail arrangement. Adjacent protofilaments are not aligned in register but are staggered about 1 nm so that the tubulin molecules form a helical array around the circumference of the microtubule. The helix is interrupted at one site where α and β subunits make lateral contacts. This produces a “seam” that runs the length of the microtubule.
749. Microtubule-Associated Proteins
750. Figure 9.10: Microtubule-associated proteins (MAPs). Schematic diagram of a brain MAP2 molecule bound to the surface of a microtubule. The MAP2 molecule shown in this figure contains three tubulin-binding sites connected by short segments of the polypeptide chain. (An alternate isoform contains four binding sites). The binding sites are spaced at a sufficient distance to allow the MAP2 molecule to attach to three separate tubulin subunits in the wall of the microtubule. The tails of the MAP molecules project outward, allowing them to interact with other cellular components.
751. Microtubules as Structural Supports and Organizers
752. Figure 9.11: Localization of microtubules of a flattened, cultured mouse cell is revealed by fluorescent anti-tubulin antibodies. Microtubules are seen to extend from the perinuclear region of the cell in a radial array. Individual microtubules can be followed and are seen to curve gradually as they conform to the shape of the cell.
753. Figure 9.12: Spatial relationship between microtubule orientation and cellulose deposition in plant cells. A wheat mesophyll cell that was double stained for microtubules (a) and cell wall cellulose (b). It is evident that the cortical microtubules and cellulose microfibrils are coaligned.
754. Microtubules as Agents of Intracellular Motility
755. Figure 9.13: Axonal transport.(a) Schematic drawing of a nerve cell showing the movement of vesicles down the length of an axon along tracks of microtubules. Vesicles move in both directions within the axon. (b) Schematic drawing of the organization of the microtubules and intermediate filaments (neurofilaments) within an axon. Vesicles containing transported materials are attached to the microtubules by cross-linking proteins, including motor proteins, such as kinesin and dynein. (c) Electron micrograph of a portion of an axon from a cultured nerve cell, showing the numerous parallel microtubules that function as tracks for axonal transport. The two membrane-bound organelles shown in this micrograph were seen under a light microscope to be moving along the axon at the time the nerve cell was fixed.
756. Axonal Transport
757. Figure 9.14: Visualizing axonal transport.(a–c) These video micrographs show the progression of a membranous organelle along a branched axon. The cell body is far out of the field to the upper left, while the termini (growth cones, page 379) are out of the field to the bottom right. The position of the organelle is indicated by the arrowheads. The organelle being followed (an autophagic vacuole) moves in the retrograde direction across the branch point and continues to move toward the cell body. Bar, 10 μm.
758. Motor Proteins that Traverse the Microtubular Cytoskeleton
759. Kinesins
760. Figure 9.15: Kinesin.(a) Structure of a kinesin-1 molecule, which consists of two heavy chains that wrap around each other to form a single, common stalk and two light chains associated with the globular ends of the heavy chains. The human genome encodes three different heavy chains and four different light chains for kinesin-1. The force-generating heads bind to the microtubule, and the tail binds to the cargo being transported. Having a molecular mass of approximately 380 kDa, kinesin is considerably smaller than the other motor proteins, myosin (muscle myosin, 520 kDa) and dynein (over 1000 kDa). (b) Schematic diagram of a kinesin molecule moving along a microtubular track. In the alternate hand-over-hand model shown here, the two heads carry out equivalent but alternating movements, not unlike that of a person walking through a garden on a linear path of stepping stones. As with walking, the trailing head (leg) moves 16 nm forward alternately on the left and right side of the stalk (body). (c) The conformational changes that occur in the head (blue) and neck (red) portions of a monomeric kinesin molecule that drive the movement of the protein along a microtubule (yellow contour map). Rather than being connected to a stalk and to a second head, the neck of this truncated kinesin molecule is attached to a GFP molecule (green). The swinging movement of the neck would normally drive the forward motion of the partner head, enabling the dimer to walk toward the plus end of a protofilament.
761. Figure 9.16: Alteration in the phenotype of a cell lacking a member of the kinesin superfamily.(a,c) Control cell from the extraembryonic tissues of a normal 9.5-day mouse embryo stained in a for microtubules (green) and in c for mitochondria (red). A significant fraction of the cell’s mitochondria are located along microtubules in the peripheral regions of the cell. (b,d) A corresponding cell obtained from an embryo that lacked both copies of the gene encoding KIF5B, which is one of three kinesin-1 isoforms in mice and humans. All of the mitochondria are clustered in the central region of the cell, suggesting that KIF5B is responsible for transporting mitochondria in an outward direction.
762. Kinesin-Mediated Organelle Transport
763. Figure 9.17: Cytoplasmic dynein and organelle transport by microtubule-tracking motor proteins.(a) Structure of a cytoplasmic dynein molecule, which contains two dynein heavy chains and a number of smaller intermediate and light chains at the base of the molecule. Each dynein heavy chain contains a large, globular, force-generating head, a protruding stalk containing a binding site for the microtubule, and a stem. (b) Schematic diagram of two vesicles moving in opposite directions along the same microtubule, one powered by kinesin moving toward the plus end of the track, and the other by cytoplasmic dynein moving toward the minus end of the track. In the model shown here, each vesicle contains both types of motor proteins, but the kinesin molecules are inactivated in the upper vesicle and the dynein molecules are inactivated in the lower vesicle. Both motor proteins are attached to the vesicle membrane by an intermediary: kinesin can be attached to vesicles by a variety of integral and peripheral membrane proteins, and dynein by a soluble protein complex called dynactin. (c) Schematic illustration of kinesin-mediated and dynein-mediated transport of vesicles, vesicular-tubular clusters (VTCs), and organelles in a nonpolarized, cultured cell.
764. Cytoplasmic Dynein
765. Figure 9.18: The centrosome.(a) Schematic diagram of a centrosome showing the paired centrioles; the surrounding pericentriolar material (PCM); and microtubules emanating from the PCM, where nucleation occurs. (b) Electron micrograph of a cross section of a centriole showing the pinwheel arrangement of the nine peripheral fibrils, each of which consists of one complete microtubule and two incomplete microtubules. (c) Electron micrograph showing two pairs of centrioles. Each pair consists of a longer parental centriole and a smaller daughter centriole (arrow), which is undergoing elongation in this phase of the cell cycle (discussed in Section 14.2). (d) Electron micrographic reconstruction of a 1.0 M potassium iodide–extracted centrosome, showing the PCM to contain a loosely organized fibrous lattice. (e) Fluorescence micrograph of a cultured mammalian cell showing the centrosome (stained yellow by an antibody against a centrosomal protein) at the center of an extensive microtubular network.
766. Microtubule-Organizing Centers (MTOCs)
767. Centrosomes
768. Figure 9.19: Microtubule nucleation at the centrosome.(a) Fluorescence micrograph of a cultured fibroblast that had been exposed to colcemid to bring about the disassembly of the cell’s microtubules and was then allowed to recover for 30 minutes before treatment with fluorescent anti-tubulin antibodies. The bright starlike structure marks the centrosome together with newly assembled microtubules that have begun to grow outward in all directions. (b) Schematic diagram of the regrowth of microtubules showing the addition of subunits at the plus end of the polymer away from the centrosome.
769. Basal Bodies and Other MTOCs
770. Figure 9.20: The role of γ-tubulin in centrosome function. (a) A dividing fibroblast that has been double stained with antibodies against γ-tubulin (red) and β-tubulin (green). Orange staining is due to the coincidence of the two types of tubulin, which occurs in the two centrosomes located at opposite poles of a dividing cell. (b) A reconstruction based on electron micrographs of a portion of a centrosome that had been incubated with purified tubulin in vitro and then labeled with antibodies against γ-tubulin. The antibodies were linked to gold particles to make them visible (as white dots) in the reconstruction. During the incubation with tubulin, the centrosome served as an MTOC to nucleate microtubules whose minus ends are seen to be labeled with clusters of gold, which are often arranged in semicircles or rings. The accompanying drawing shows the outline of the microtubule seen in the reconstruction. (c) A model for γ-tubulin function during the assembly of microtubules. Nucleation begins as αβ-tubulin dimers bind to an open ring of γ-tubulin molecules (brown), which are held in place by a number of non-tubulin proteins (green) that make up the large γ-TuRC. Nucleation by the γ-TuRC defines microtubule polarity with a ring of α-tubulin monomers situated at the minus end of the structure.
771. Microtubule Nucleation
772. The Dynamic Properties of Microtubules
773. Figure 9.21: Four major arrays of microtubules present during the cell cycle of a plant cell. The organization of the microtubules at each stage are described in the text.
774. Figure 9.22: Nucleation of plant cortical microtubules. (a) The micrographs show a portion of a live cultured tobacco cell that is expressing fluorescently labeled GFP-tubulin. During the period of observation, an existing microtubule of the cortex nucleates the assembly of a daughter microtubule, which grows outward forming a Y-shaped branch. The end of a newly formed microtubule is indicated by the arrowhead, the branchpoint by the arrow. (b) Electron micrograph of a microtubule with two daughter microtubules branching from its surface. The branched microtubules were assembled in a cell-free system containing tubulin subunits. Bar, 10 μm. (c) A schematic model showing how new microtubules are nucleated at the sites of γ-tubulin present on the surface of an existing microtubule.
775. The Underlying Basis of Microtubule Dynamics
776. Figure 9.23: Microtubules assembled in the test tube. Electron micrograph of frozen, unfixed microtubules that had polymerized in vitro. The individual protofilaments and their globular tubulin subunits are visible. Note that the middle of the three microtubules contains only 11 protofilaments.
777. Figure 9.24: The assembly of tubulin onto an existing microtubular structure. Electron micrograph showing the in vitro assembly of brain tubulin onto the plus ends of the microtubules of an axoneme from a Chlamydomonas flagellum.
778. Figure 9.25: The structural cap model of dynamic instability. According to the model, the growth or shrinkage of a microtubule depends on the state of the tubulin dimers at the plus end of the microtubule. Tubulin-GTP dimers are depicted in red. Tubulin-GDP dimers are depicted in blue. In a growing microtubule (step 1), the tip consists of an open sheet containing tubulin-GTP subunits. In step 2, the tube has begun to close, forcing the hydrolysis of the bound GTP. In step 3, the tube has closed to its end, leaving only tubulin-GDP subunits. GDP-tubulin subunits have a curved conformation compared to their GTP-bound counterparts, which makes them less able to fit into a straight protofilament. The strain resulting from the presence of GDP-tubulin subunits at the plus end of the microtubule is released as the protofilaments curl outward from the tubule and undergo catastrophic shrinkage (step 4). (b) Cryo-EM image of the growing end of a microtubule showing a curved, open sheet. (c) Cryo-EM image of the shrinking end of a microtubule showing outwardly curved protofilaments.
779. Figure 9.26: Microtubule dynamics in living cells. This cultured fibroblast was injected with a small volume of tubulin that had been covalently linked to biotin, a small molecule whose location in the cell is readily determined using fluorescent anti-biotin antibodies. Approximately one minute following injection, the cells were fixed, and the location of biotinylated tubulin that had been incorporated into insoluble microtubules was determined. It is evident from this fluorescence micrograph that, even during periods as short as one minute, tubulin subunits are widely incorporated at the growing ends of cytoskeletal microtubules.
780. Figure 9.27: Dynamic instability. This series of photographs shows the changes in length of a single microtubule in the growth cone of a neuron. The cell was microinjected with fluorescently labeled tubulin at sufficiently low concentration to produce green fluorescent speckles along the length of microtubules. As discussed on page 327, such speckles provide fixed reference points that can be followed over time. Each of the yellow horizontal lines connects one of these speckles from one time point to another. The blue line indicates the approximate plus end of the microtubule at various time points. From 0 to about 200 seconds, the microtubule undergoes a gradual addition of tubulin subunits at its plus end. Then, from about 200 to 240 seconds the microtubule experiences rapid shrinkage.
781. Figure 9.28: Binding of a microtubule plus-end tracking protein (+TIP). The micrograph shows a live image of a human lung fibroblast expressing mCherry-labeled tubulin (generating red microtubules) and GFP-labeled EB3, a +TIP (appearing green). EB3 is seen to bind to the plus ends of the microtubules.
782. Cilia and Flagella: Structure and Function
783. Figure 9.29: Ciliary beat. (a) The various stages in the beat of a cilium. (b) The cilia on the surface of a ciliated protozoan beat in metachronal waves in which the cilia in a given row are in the same stage of the beat cycle, but those in adjacent rows are in a different stage. RS, cilia in recovery stroke; ES, cilia in effective power stroke. (c) Human bronchial epithelial cells in which the cilia at the luminal surface are labeled red and the nuclei are labeled blue.
784. Figure 9.30: Eukaryotic flagella. (a) The unicellular alga Chlamydomonas reinhardtii. The two flagella appear green after binding a fluorescent antibody directed against a major flagellar membrane protein. The red color results from autofluorescence of the cell’s chlorophyll. Unlike many flagellated organisms, Chlamydomonas does not need its flagella to survive and reproduce, so that mutant strains exhibiting various types of flagellar defects can be cultured. (b) The forward movement of Chlamydomonas is accomplished by an asymmetric waveform that resembles the breast stroke. A different type of flagellar waveform is shown in Figure 9.35.
785. Figure 9.31: The structure of a ciliary or flagellar axoneme. (a) Cross section through a sperm axoneme. The peripheral doublets are seen to consist of a complete and an incomplete microtubule, whereas the two central microtubules are complete. The dynein arms are seen as “fuzzy” projections from the wall of the complete microtubule. The molecular structure of these arms is discussed in a later section. (b) Schematic diagram of an axoneme of a protist showing the structure of the microtubular fibers, the two types of dynein arms (three-headed outer arms and two-headed inner arms), the nexin links between the doublets, the central sheath surrounding the central microtubules, and the radial spokes projecting from the outer doublets toward the central sheath. A more detailed and complex view of the molecular architecture of the axoneme has been obtained with the application of cryoelectron tomography (Section 18.2; see PNAS 102:15889, 2005, Science 313:944, 2006, and J. Cell Biol. 187: 921, 2009. (Note: The cilia and flagella of animals typically contain two-headed outer dynein arms.)
786. Figure 9.32: Longitudinal view of an axoneme. (a) Electron micrograph of a median longitudinal section of a straight region of a cilium. The radial spokes are seen joining the central sheath with the A microtubule of the doublet. (b) Schematic diagram of a longitudinal section of a flagellar doublet. Radial spokes emerge in groups of three, which repeat (at 96 nm in this case) along the length of the microtubule. Outer dynein arms are spaced at intervals of 24 nm.
787. Figure 9.33: Basal bodies and axonemes. (a) Electron micrograph of a longitudinal section through the basal bodies of a number of cilia at the apical surface of epithelial cells of a rabbit oviduct. These basal bodies arise from centrioles that are generated in the cytoplasm and migrate to sites beneath the plasma membrane. (b) Schematic diagram of the structural relationship between the microtubules of the basal body and the axoneme of a cilium or flagellum.
788. THE HUMAN PERSPECTIVE: The Role of Cilia in Development and Disease
789. Figure 1: Primary cilia. Immunofluorescence image showing a single primary cilium (green) projecting from the apical surface of each epithelial cell of the collecting duct of the kidney. The intercellular junctions between the epithelial cells are shown in red.
790. Intraflagellar Transport
791. Figure 9.34: Intraflagellar transport (IFT). Electron micrograph of a longitudinal section of a Chlamydomonas flagellum showing two rows of protein particles (bounded by arrowheads) situated between the outer doublet microtubules and the flagellar plasma membrane. As illustrated in the insets, each row of particles and its associated cargo of axonemal proteins are moved along the outer doublet microtubule by a motor protein, either cytoplasmic dynein 2 if they are moving toward the flagellar base or kinesin 2 if the particles are moving toward the tip of the flagellum.
792. The Dynein Arms
793. Figure 9.35: A sea urchin sperm reactivated with 0.2 mM ATP following demembranation with the detergent Triton X-100. This multiple-exposure photomicrograph was obtained with five flashes of light and shows the reactivated flagellum at different stages of its beat.
794. Figure 9.36: Steps in the chemical dissection of cilia from the protozoan Tetrahymena. The numbered steps are described in the text.
795. The Mechanism of Ciliary and Flagellar Locomotion
796. Figure 9.37: A model of the structure and function of flagellar/ciliary dynein. (a) Platinum replica of a rotary-shadowed, flagellar outer-arm dynein molecule prepared by rapid-freeze, deep-etch electron microscopy. Each of the three heavy chains forms a prominent globular head with an extension (stalk) that functions in linking the dynein arm to the neighboring doublet. An interpretive drawing is shown on the right. (b–d) High-resolution micrographs (with b′, c′ interpretive models below) of a flagellar dynein heavy chain before (b) and after (c) its power stroke. The motor domain, which consists of a number of modules arranged in a wheel, is seen to have rotated, which has caused the stalk to move in a leftward direction. The image shown in (d) is a composite of molecules showing the position of the stalk before and after the power stroke. The power stroke causes the microtubule bound to the tip of the stalk to slide 15 nm in a leftward direction relative to the motor domain. In vitro motility assays suggest that dynein may be capable of “shifting gears” so that it can take shorter, more powerful steps as it moves a load of increasing mass. (Note: An alternate model of dynein function is discussed in Science 322:1647, 2008.)
797. Figure 9.38: Schematic representation of the forces that drive ciliary or flagellar motility. The steps are described in the text. A true representation of the power stroke is shown in the previous figure.
798. Figure 9.39: The sliding-microtubule mechanism of ciliary or flagellar motility. Schematic diagram of the sliding of neighboring microtubules relative to one another. When the cilium is straight, all the outer doublets end at the same level (center). Cilium bending occurs when the doublets on the inner side of the bend slide beyond those on the outer side (top and bottom). The movement of the dynein arms responsible for the sliding of neighboring microtubules was shown in the previous figures.
799. REVIEW
800. Figure 9.40: Experimental demonstration of microtubule sliding. Sea urchin sperm were demembranated, reactivated with ATP, and photographed by multiple exposure as in Figure 9.35. In this experiment, gold beads were attached to the exposed outer doublet microtubules, where they could serve as markers for specific sites along different doublets. As the flagellum underwent beating, the relative positions of the beads were monitored. As shown here, the beads moved farther apart and then closer together as the flagellum undulated, indicating that the doublets are sliding back and forth relative to one another.
801. 9.4: Intermediate Filaments
802. Table 9.2: Properties and Distribution of the Major Mammalian Intermediate Filament Proteins
803. Figure 9.41: Cytoskeletal elements are connected to one another by protein cross-bridges. Electron micrograph of a replica of a small portion of the cytoskeleton of a fibroblast after selective removal of actin filaments. Individual components have been digitally colorized to assist visualization. Intermediate filaments (blue) are seen to be connected to microtubules (red) by long wispy cross-bridges consisting of the fibrous protein plectin (green). Plectin is localized by antibodies linked to colloidal gold particles (yellow).
804. Intermediate Filament Assembly and Disassembly
805. Figure 9.42: A model of intermediate filament assembly and architecture. Each monomer has a pair of globular terminal domains (red or yellow) separated by a long α-helical region (step 1). Pairs of monomers associate in parallel orientation with their ends aligned to form dimers (step 2). Depending on the type of intermediate filament, the dimers may be composed of identical monomers (homodimers) or nonidentical monomers (heterodimers). Dimers in turn associate in an antiparallel, staggered fashion to form tetramers (step 3), which are thought to be the basic subunit in the assembly of intermediate filaments. In the model shown here, 8 tetramers associate laterally to form a unit length of the intermediate filament (step 4). Highly elongated intermediate filaments are then formed from the end-to-end association of these unit lengths (step 5). Once formed, intermediate filaments undergo a process of dynamic remodeling that is thought to involve the intercalation of unit lengths of filament into the body of an existing filament (step 6). (b) A model of a tetramer of the IF protein vimentin.
806. Figure 9.43: Experimental demonstration of the dynamic character of intermediate filaments. These photographs show the results of an experiment in which biotin-labeled type I keratin was microinjected into cultured epithelial cells and localized 20 minutes later using immunofluorescence. The photograph in a shows the localization of the injected biotinylated keratin (as revealed by fluorescent anti-biotin antibodies) that had become incorporated into filaments during the 20-minute period following injection. The photograph in b shows the distribution of intermediate filaments in the cell as revealed by anti-keratin antibodies. The dotlike pattern of fluorescence in a indicates that the injected subunits are incorporated into the existing filaments at sites throughout their length, rather than at their ends. (Compare with a similar experiment with labeled tubulin in Figure 9.26.) Bar, 10 μm.
807. Types and Functions of Intermediate Filaments
808. REVIEW
809. 9.5: Microfilaments
810. Figure 9.44: The organization of intermediate filaments (IFs) within an epithelial cell. (a) In this schematic drawing, IFs are seen to radiate throughout the cell, being anchored at both the outer surface of the nucleus and the inner surface of the plasma membrane. Connections to the nucleus are made via proteins that span both membranes of the nuclear envelope and to the plasma membrane via specialized sites of adhesion such as desmosomes and hemidesmosomes. IFs are also seen to be interconnected to both of the other types of cytoskeletal fibers. Connections to microtubules (MTs) and microfilaments (MFs) are made primarily by members of the plakin family of proteins, such as the dimeric plectin molecule shown in Figure 9.41. (b) Distribution of keratin-containing intermediate filaments in cultured skin cells (keratinocytes). The filaments are seen to form a cagelike network around the nucleus and also extend to the cell periphery
811. Figure 9.45: Actin filament structure.(a) A model of an actin filament. The actin subunits are represented in three colors to distinguish the consecutive subunits more easily. The subdomains in one of the actin subunits are labeled 1, 2, 3, and 4 and the ATP-binding cleft in each subunit is evident. Actin filaments have polarity, which is denoted as a plus and minus end. The cleft (in the upper red subunit) is present at the minus end of the filament. (b) Electron micrograph of a replica of an actin filament showing its double-helical architecture.
812. Microfilament Assembly and Disassembly
813. Figure 9.46: Determining the location and polarity of actin filaments using the S1 subunit of myosin. Electron micrograph of a replica showing the microfilament bundles in the core of the microvilli of an intestinal epithelial cell. The cell had been fixed, treated with S1 myosin fragments, freeze fractured, and deep etched to expose the filamentous components of the cytoplasm. The intermediate filaments (IFs) at the bottom of the micrograph do not contain actin and therefore do not bind the S1 myosin fragments. These intermediate filaments originate at the desmosomes of the lateral surfaces of the cell.
814. Figure 9.47: Actin assembly in vitro.(a) Electron micrograph of a short actin filament that was labeled with S1 myosin and then used to nucleate actin polymerization. The addition of actin subunits occurs much more rapidly at the barbed (plus) end than at the pointed (minus) end of the existing filament. (b) Schematic diagram of the kinetics of actin-filament assembly in vitro. All of the orange subunits are part of the original seed; red subunits were present in solution at the beginning of the incubation. The steps are described in the text. Once a steady-state concentration of monomers is reached, subunits are added to the plus end at the same rate they are released from the minus end. As a result, subunits treadmill through the filament in vitro. Note: No attempt is made to distinguish between subunits with a bound ATP versus ADP.
815. Myosin: The Molecular Motor of Actin Filaments
816. Conventional (Type II) Myosins
817. Figure 9.48: Experimental demonstration of a role for myosin II in the directional movement of growth cones.(a) Fluorescence micrograph showing fine processes (neurites) growing out from a microscopic fragment of mouse embryonic nervous tissue. The neurites (stained green) are growing outward on a glass coverslip coated with strips of laminin (stained red). Laminin is a common component of the extracellular matrix (page 243). The tip of each nerve process contains a motile growth cone. When the growth cones reach the border of the laminin strip (indicated by the line with arrowheads), they turn sharply and continue to grow over the laminin-coated surface. Bar, 500 μm. (b) The tissue in this micrograph was obtained from a mouse embryo lacking myosin IIB. The growth cones no longer turn when they reach the edge of the laminin-coated surface, causing the neurites to grow forward onto a surface (black) lacking laminin. Bar, 80 μm.
818. Figure 9.49: Structure of a myosin II molecule.(a) Electron micrograph of negatively stained myosin molecules. The two heads and tail of each molecule are clearly visible. (b) A highly schematic drawing of a myosin II molecule (molecular mass of 520,000 daltons). The molecule consists of one pair of heavy chains (purple) and two pairs of light chains, which are named as indicated. The paired heavy chains consist of a rod-shaped tail in which portions of the two polypeptide chains wrap around one another to form a coiled coil and a pair of globular heads. When treated with a protease under mild conditions, the molecule is cleaved at the junction between the neck and tail, which generates the S1 fragment.
819. Figure 9.50: In vitro motility assay for myosin.(a) Schematic drawing in which myosin heads are bound to a silicone-coated coverslip, which is then incubated with a preparation of actin filaments. (b) Results of the experiment depicted in a. Two images were taken 1.5 seconds apart and photographed as a double exposure on the same frame of film. The dashed lines with arrowheads show the sliding movement of the actin filaments over the myosin heads during the brief period between exposures.
820. Figure 9.51: Structure of a bipolar myosin II filament.(a) Schematic diagram of the staggered arrangement of the individual myosin molecules in a myosin II filament. (b) Electron micrograph of a bipolar myosin filament formed in vitro. The heads of the filament are seen at both ends, leaving a smooth section in the center of the filament.
821. Unconventional Myosins
822. Figure 9.52: Myosin V—a two-headed unconventional myosin involved in organelle transport.(a) Direct visualization of a single myosin V molecule (one that is lacking its normal tail domain) as it moves along an actin filament in vitro in the presence of ATP. Using high-speed atomic force microscopy (HS-AFM), this series of images shows the movement of the molecule over a period of about one second. (b) Succesive HS-AFM images showing the hand-over-hand movement of a single myosin V molecule as it passes through a cluster of obstacles (consisting of streptavidin protein molecules). The swinging neck (or lever arm) is highlighted with a thin white line. Interpretive drawings (b′) depict the motor protein as seen in several of the corresponding HS-AFM images. A continuous movie of the protein’s movement can be seen in the supplement to this article. (c) Schematic drawing of a complete dimeric myosin V molecule, including its numerous light chains, with both heads bound to an actin filament and its tail domain bound to a vesicle. Rab27a and melanophilin serve as adaptors that link the globular ends of the tail to the vesicle membrane. The long neck of myosin V binds 6 light chains.
823. Figure 9.53: The contrasting roles of microtubule- and microfilament-based motors in intracellular transport. Most vesicle transport is thought to be mediated by members of the kinesin and dynein families, which carry their cargo over relatively long distances. It is thought that some vesicles also carry myosin motors, such as myosin Va, which transport their cargo over microfilaments, including those present in the peripheral (cortical) regions of the cell. The two types of motors may act in a cooperative manner, as illustrated here in the case of a frog pigment cell in which pigment granules move back and forth within extended cellular processes.
824. REVIEW
825. 9.6: Muscle Contractility
826. Figure 9.54: Hair cells, actin bundles, and unconventional myosins. (a) Drawing of a hair cell of the cochlea. The inset shows a portion of several of the stereocilia, which are composed of a tightly grouped bundle of actin filaments. (b) Transmission electron micrograph of a cross section of a stereocilium showing it is composed of a dense bundle of actin filaments. (c) Fluorescence micrograph of a hair cell from the vestibule of a rat inner ear. The tips of the stereocilia are labeled green due to the incorporation of GFP-actin monomers at their barbed ends. Taller stereocilia contain a longer column of GFP-labeled subunits, which reflects a more rapid incorporation of actin monomers. The stereocilia appear red due to labeling by rhodamine-labeled phalloidin, which binds to actin filaments. (d) A hair cell from the bullfrog inner ear. The localization of myosin VIIa is indicated in green. The orange bands near the bases of the stereocilia (due to red and green overlap) indicate a concentration of myosin VIIa. (e) Myosin XVa (green) is localized at the tips of the stereocilia of a rat auditory hair cell. (f) Scanning electron micrograph of the hair cells of the cochlea of a control mouse. The stereocilia are arranged in V-shaped rows. (g) A corresponding micrograph of the hair cells of a mouse with mutations in the gene encoding myosin VIIa, which causes deafness. The stereocilia of the hair cells exhibit a disorganized arrangement.
827. Figure 9.55: The structure of skeletal muscle.(a)Levels of organization of a skeletal muscle. (b) Light micrograph of a multinucleated muscle fiber. (c) Electron micrograph of a sarcomere with the bands lettered.
828. The Sliding Filament Model of Muscle Contraction
829. Figure 9.56: The contractile machinery of a sarcomere.(a) Diagram of a sarcomere showing the overlapping array of thin actin-containing (orange) and thick myosin-containing (purple) filaments. The small transverse projections on the myosin fiber represent the myosin heads (cross-bridges). (b) Electron micrograph of a cross section through an insect flight muscle showing the hexagonal arrangement of the thin filaments around each thick filament.
830. Figure 9.57: The shortening of the sarcomere during muscle contraction.(a) Schematic diagram showing the difference in the structure of the sarcomere in a relaxed and contracted muscle. During contraction, the myosin cross-bridges make contact with the surrounding thin filaments, and the thin filaments are forced to slide toward the center of the sarcomere. Cross-bridges work asynchronously, so that only a fraction are active at any given instant. (b) Electron micrographs of longitudinal sections through a relaxed (top) and contracted (bottom) sarcomere. The micrographs show the disappearance of the H zone as the result of the sliding of the thin filaments toward the center of the sarcomere.
831. The Composition and Organization of Thick and Thin Filaments
832. Figure 9.58: The molecular organization of the thin filaments. Each thin filament consists of a helical array of actin subunits with rod-shaped tropomyosin molecules situated in the grooves and troponin molecules spaced at defined intervals, as described in the text. The positional changes in these proteins that trigger contraction are shown in Figure 9.63.
833. Figure 9.59: The arrangement of titin molecules within the sarcomere. These huge elastic molecules stretch from the end of the sarcomere at the Z line to the M band in the sarcomere center. Titin molecules are thought to maintain the thick filaments in the center of the sarcomere during contraction. The I-band portion of the titin molecule contains spring-like domains and is capable of great elasticity. The nebulin molecules (which are not discussed in the text) are thought to act like a “molecular ruler” by regulating the number of actin monomers that are allowed to assemble into a thin filament.
834. The Molecular Basis of Contraction
835. Figure 9.60: Model of the swinging lever arm of a myosin II molecule.(a) During the power stroke, the neck of the myosin molecule moves through a rotation of approximately 70°, which would produce a movement of the actin filament of approximately 10 nm. (b) A model of the power stroke of the myosin motor domain consisting of the motor domain (or head) and adjoining neck (or lever arm). An attached actin filament is shown in gray/brown. The long helical neck is shown in two positions, both before and after the power stroke (depicted as the upper dark blue and lower light blue necks, respectively). It is this displacement of the neck region that is thought to power muscle movement. The essential and regulatory light chains, which wrap around the neck, are shown in yellow and magenta, respectively.
836. The Energetics of Filament Sliding
837. Figure 9.61: Video: A schematic model of the actinomyosin contractile cycle. The movement of the thin filament by the force-generating myosin head occurs as the result of a linkage between a mechanical cycle involving the attachment, movement, and detachment of the head and a chemical cycle involving the binding, hydrolysis, and release of ATP, ADP, and Pi. In this model, the two cycles begin in step 1 with the binding of ATP to a cleft in the myosin head, causing the detachment of the head from the actin filament. The hydrolysis of the bound ATP (step 2) energizes the head, causing it to bind weakly to the actin filament (step 3). The release of Pi causes a tighter attachment of the myosin head to the thin filament and the power stroke (step 4) that moves the thin filament toward the center of the sarcomere. The release of the ADP (step 5) sets the stage for another cycle.
838. Excitation–Contraction Coupling
839. Figure 9.62: The functional anatomy of a muscle fiber. Calcium is housed in the elaborate network of internal membranes that make up the sarcoplasmic reticulum (SR). When an impulse arrives by means of a motor neuron, it is carried into the interior of the fiber along the membrane of the transverse tubule to the SR. The calcium gates of the SR open, releasing calcium into the cytosol. The binding of calcium ions to troponin molecules of the thin filaments leads to the events described in the following figure and the contraction of the fiber.
840. Figure 9.63: The role of tropomyosin in muscle contraction. Schematic diagram of the steric hindrance model in which the myosin-binding site on the thin actin filaments is controlled by the position of the tropomyosin molecule. When calcium levels rise, the interaction between calcium and troponin (not shown) leads to a movement of the tropomyosin from position b to position a, which exposes the myosin-binding site on the thin filament to the myosin head.
841. REVIEW
842. 9.7: Nonmuscle Motility
843. Actin-Binding Proteins
844. Figure 9.64: Two different arrangements of actin filaments within a cell. As described later in the chapter, cells move across a substratum by extending various types of processes. This electron micrograph of the leading edge of a motile fibroblast shows the high density of actin filaments. These filaments are seen to be organized into two distinct arrays: as bundles in which the filaments are arranged parallel to one another (arrow) and as a cross-linked meshwork in which the filaments are arranged in various directions.
845. Figure 9.65: The roles of actin-binding proteins.
846. Figure 9.66: Actin filaments and actin-binding proteins in a microvillus. Microvilli are present on the apical surface of epithelia that function in absorption of solutes, such as the lining of the intestine and wall of the kidney tubule. Actin filaments are maintained in a highly ordered arrangement by the bundling proteins villin and fimbrin. The role of myosin I, which is present between the plasma membrane of the microvillus and the peripheral actin filaments, remains unclear.
847. Examples of Nonmuscle Motility and Contractility
848. Actin Polymerization as a Force-Generating Mechanism
849. Cell Locomotion
850. Figure 9.67: Cell motility can be driven by actin polymerization.(a) Fluorescence micrograph of a portion of a cell infected with the bacterium L. monocytogenes. The bacteria appear as red-stained objects just in front of the green-stained filamentous actin tails. (b) Electron micrograph of a cell infected with the same bacterium as in a, showing the actin filaments that form behind the bacterial cell and push it through the cytoplasm. The actin filaments have a bristly appearance because they have been decorated with myosin heads. Bar at the upper left, 0.1 μm.
851. Figure 9.68: Scanning electron micrograph of a mouse fibroblast crawling over the surface of a culture dish. The leading edge of the cell is spread into a flattened lamellipodium whose structure and function are discussed later in the chapter.
852. Figure 9.69: The repetitive sequence of activities that occurs as a cell crawls over the substratum. Step 1 shows the protrusion of the leading edge of the cell in the form of a lamellipodium. Step 2 shows the adhesion of the lower surface of the lamellipodium to the substratum, an attachment that is mediated by integrins residing in the plasma membrane. The cell uses this attachment to grip the substratum. Step 3 shows the movement of the bulk of the cell forward over the site of attachment, which remains relatively stationary. This movement is accomplished by a contractile (traction) force exerted against the substratum. Step 4 shows the cell after the rear attachments with the substratum have been severed and the trailing portion of the cell has been pulled forward.
853. Cells that Crawl over the Substratum
854. Figure 9.70: The leading edge of a motile cell.(a)The leading edge of this motile fibroblast is flattened against the substratum and spread out into a veil-like lamellipodium. (b) Scanning electron micrograph of the leading edge of a cultured cell, showing the ruffled membranes of the lamellipodium.
855. Figure 9.71: Directed cell motility.
856. Figure 9.72: The structural basis of lamellipodial extension. Electron micrograph of a replica of the cytoskeleton at the leading edge of a motile mouse fibroblast. The actin filaments are seen to be arranged in a branched network, which has been colorized to indicate individual “trees.” The circular insets show a succession of Y-shaped junctions between branched actin filaments. Arp2/3 complexes are localized at the base of each branch by antibodies linked to colloidal gold particles (yellow).
857. Figure 9.73: Distribution of traction forces within a migrating fibroblast.(a) As a cell migrates it generates traction (pulling) forces against its substrate. The present image shows the traction forces generated per unit area by the surface of a migrating fibroblast. Traction forces were calculated at different sites on the surface based on the degree of substrate deformation (see Figure 7.18). The magnitude of the traction forces are expressed by varying colors with red representing the strongest forces. The largest forces are generated at sites of small focal complexes that form transiently behind the leading edge of the cell where the lamellipodium is being extended (arrow). Deformation at the rear of the cell (shown in red) occurs as the front end actively pulls against the tail, which is passively anchored. (b) A living, migrating fibroblast exhibiting a well-developed lamellipodium that is adhering to the underlying substratum at numerous sites (red). This cell is expressing GFP-actin (green) and had been injected with rhodamine-tagged vinculin (red). The fluorescently labeled vinculin is incorporated into dot-like focal complexes near the leading edge of the cell. Some of these focal complexes disassemble, whereas others mature into focal adhesions, which are situated farther from the advancing edge.
858. Axonal Outgrowth
859. Figure 9.74: The roles of actin and myosin in the lamellipodial-based movement of fish keratocytes.(a,b) Fluorescence micrographs of a fish keratocyte moving over a culture dish by means of a broad, flattened lamellipodium. The arrow shows the direction of movement, which can occur at rates of 10 μm/min. The distribution of filamentous actin is revealed in part a, which shows the localization of fluorescently labeled phalloidin, which binds only to actin filaments. The distribution of myosin in the same cell is revealed in part b, which shows the localization of fluorescent antimyosin antibodies. It is evident that the body of the lamellipodium contains actin filaments but is virtually devoid of myosin. Myosin is concentrated, instead, in a band that lies just behind the lamellipodium, where it merges with the body of the cell. (c) A schematic drawing depicting the filamentous actin network of the lamellipodium and the actin–myosin interactions toward the rear of the lamellipodium. The actin network is shown in red, myosin molecules in blue.
860. Figure 9.75: The structure of a growth cone: the motile tip of a growing axon.(a) A video image of a live growth cone. The terminus is spread into a flattened lamellipodium that creeps forward over the substratum. Rodlike microspikes (arrows) can be seen within the transparent veil of the lamellipodium, and fine processes called filopodia (arrowheads) can be seen projecting ahead of the leading edge of the lamellipodium. Bar, 5 μm. (b) Fluorescence micrograph of the growth cone of a neuron showing the actin filaments (green) concentrated in the peripheral domain and the microtubules (orange) concentrated in the central domain. A number of microtubules can be seen to invade the peripheral domain, where they interact with actin-filament bundles.
861. Figure 9.76: The directed movements of a growth cone.(a) A video image of a live growth cone of a Xenopus neuron that has turned toward a diffusible protein (netrin-1) released from a pipette whose position is indicated by the arrow. (b) The growth cone (green) at the tip of a motor axon has made contact by means of its filopodia with a target cell that is expressing the neuronal guidance factor ephrin (red).
862. Changes in Cell Shape during Embryonic Development
863. REVIEW
864. Figure 9.77: Early stages in the development of the vertebrate nervous system.(a–d) Schematic drawings of the changes in cell shape that cause a layer of flattened ectodermal cells at the mid-dorsal region of the embryo to roll into a neural tube. The initial change in height of the cells is thought to be driven by the orientation and elongation of microtubules, whereas the rolling of the plate into a tube is thought to be driven by contractile forces generated by actin filaments at the apical ends of the cells. (e) Scanning electron micrograph of the dorsal surface of a chick embryo as its neural plate is being folded to form a tube.
865. Synopsis
866. Analytic Questions
867. 10: The Nature of the Gene and the Genome
868. Model of DNA built by James Watson and Francis Crick at Cambridge University, 1953. Inset shows a photograph taken by Rosalind Franklin of the X-ray diffraction pattern of a DNA fiber that suggested the helical nature of DNA.
869. Figure 10.1: An overview depicting several of the most important early discoveries on the nature of the gene. Each of these discoveries is discussed in the present chapter.
870. 10.1: The Concept of a Gene as a Unit of Inheritance
871. Table 10.1: Seven Traits of Mendel’s Pea Plants
872. 10.2: Chromosomes: The Physical Carriers of the Genes
873. The Discovery of Chromosomes
874. Figure 10.2: Events occurring in the roundworm Ascaris following fertilization, as reported in a classic nineteenth-century investigation. Both the male and female gamete are seen to contain two chromosomes. Fusion of the sperm and egg nuclei (called pronuclei) in the egg cytoplasm (between e and f) produces a zygote containing four chromosomes. The second polar body shown in a is a product of the previous meiosis as described in Section 14.3.
875. Chromosomes as the Carriers of Genetic Information
876. Figure 10.3: Homologous chromosomes. Sutton’s drawing of the homologous chromosomes of the male grasshopper that have associated during meiotic prophase to form bivalents. Eleven pairs of homologous chromosomes (a–k) and an unpaired X chromosome were observed.
877. The Chromosome as a Linkage Group
878. Genetic Analysis in Drosophila
879. Figure 10.4: The fruit fly Drosophila melanogaster. Photograph of a wild-type female fruit fly and a mutant male that carries a mutation leading to white eyes.
880. Crossing Over and Recombination
881. Figure 10.5: Fruit flies have four pairs of homologous chromosomes, one of which is very small. The two dissimilar homologues are the chromosomes that determine sex. As in humans, male fruit flies are XY and females are XX.
882. Figure 10.6: Visualizing sites of crossing over. Homologous chromosomes wrap around each other during meiosis, as seen in this micrograph of a meiotic cell of a lily. The points at which the homologues are crossed are termed chiasmata (arrows) and (as discussed in Chapter 14) are sites at which crossing over had occurred at an earlier stage.
883. Figure 10.7: Crossing over provides the mechanism for reshuffling alleles between maternal and paternal chromosomes.
884. Mutagenesis and Giant Chromosomes
885. Figure 10.8: Giant polytene chromosomes of larval insects. (a) These giant polytene chromosomes from the salivary gland of a larval fruit fly show several thousand distinct, darkly staining bands. The bands have been identified as the loci of particular genes. The inset shows how polytene chromosomes consist of a number of individual DNA molecules. The stained bands on the chromosomes correspond to sites where the DNA is more tightly compacted. (b) Scanning electron micrograph of a giant polytene chromosome from a Chironomus larva showing how specific sites are expanded to form a “puff.” Chromosome puffs are sites where DNA is being very actively transcribed.
886. REVIEW
887. 10.3: The Chemical Nature of the Gene
888. The Structure of DNA
889. Figure 10.9: The chemical structure of DNA. (a) Model of a DNA nucleotide containing the base thymine; the molecule is deoxythymidine 5′-monophosphate (dTMP). The netlike cage represents the electron density of the atoms that make up the molecule. (b) Chemical structure of a DNA nucleotide containing the base adenosine; the molecule is deoxyadenosine 5′-monophosphate (dAMP). A nucleotide is composed of a nucleoside linked to a phosphate; the nucleoside portion of the molecule (i.e., deoxyadenosine) is enclosed by the dashed line. (c) The chemical structure of a small segment of a single DNA strand showing all four nucleotides.
890. Base Composition
891. The Watson-Crick Proposal
892. Figure 10.10: The double helix. (opposite) (a) Schematic representation of the DNA double helix. (b) Space-filling model of the B form of DNA. (c) The Watson-Crick base pairs. The original model showed both A-T and G-C pairs with two hydrogen bonds; the third hydrogen bond in the G-C pair was subsequently identified by Linus Pauling. (d) Electron micrograph of DNA being released from the head of a T2 bacteriophage. This linear DNA molecule (note the two free ends) measures 68 μm in length, approximately 60 times longer than the phage head in which it is contained.
893. The Importance of the Watson-Crick Proposal
894. Figure 10.11: Three functions required of the genetic material. (a) DNA must contain the information that encodes inheritable traits. (b) DNA must contain the information that directs its own duplication. (c) DNA must contain the information that directs the assembly of specific proteins.
895. DNA Supercoiling
896. Figure 10.12: Supercoiled DNA. (a,b) Electron micrographs showing the differences in conformation between a relaxed, circular molecule of phage DNA (a) and the same type of molecule in a supercoiled state (b). (c) When a mixture of relaxed and supercoiled SV40 DNA molecules are subjected to gel electrophoresis, the highly compact, supercoiled form (seen at the bottom of the gel) moves much more rapidly than the relaxed form. The DNA molecules are visualized by staining the gel with ethidium bromide, a fluorescent molecule that inserts itself into the double helix.
897. Figure 10.13: Underwound DNA. The DNA molecule at the left is underwound; that is, it has more than an average of 10 base pairs per turn of a helix. An underwound molecule spontaneously assumes a negatively supercoiled conformation, as shown on the right.
898. REVIEW
899. 10.4: The Structure of the Genome
900. Figure 10.14: DNA topoisomerases. (a) A model depicting the action of human topoisomerase I. The enzyme (yellow) cuts one of the strands of the DNA (step 1), which rotates around a phosphodiester bond in the intact strand. The cut strand is then resealed (step 2). (Note: The drawing depicts a type IB topoisomerase; type IA enzymes found in bacteria act by a different mechanism.) (b) A molecular model based on X-ray crystallography depicting the action of topoisomerase II, a dimeric enzyme consisting of two identical halves. In step 1, the enzyme has bound the G-DNA segment, so named because it will form the gate through which the T-DNA (or transported DNA) segment will pass. In step 2, the dimeric enzyme has hydrolyzed two molecules of ATP and undergone a conformational change as the two ATPase domains close. In step 3, the G-segment is cleaved, and the T-segment is passed through the open “gate.” At this stage, both cut ends of the G-segment are covalently bound to the enzyme. In step 4, the two ends of the G-segment are rejoined, and the T-segment exits via the C gate. (c) Types of reactions that are catalyzed by topoisomerases. Part 1 shows supercoiling–relaxation reactions; part 2 shows knotting–unknotting reactions; part 3 shows catenation–decatenation reactions. (d) Electron micrograph of a pair of interconnected (catenated) circular DNA molecules. Molecules of this type collect in bacteria lacking a specific topoisomerase.
901. The Complexity of the Genome
902. DNA Denaturation
903. Figure 10.15: Thermal denaturation of DNA. A thermal denaturation curve for native bacteriophage T6 DNA in 0.3 M sodium citrate. The “melting” of the DNA (strand separation) occurs over a narrow range of temperature, particularly for the simpler DNAs of small viruses. The temperature corresponding to half the increase in absorbance is termed the (Tm)
904. DNA Renaturation
905. The Complexity of Viral and Bacterial Genomes
906. The Complexity of Eukaryotic Genomes
907. Figure 10.16: The kinetics of renaturation of viral and bacterial DNAs. The curves show the renaturation of sheared strands of DNA from two viruses (MS-2 and T4) and a bacterium (E. coli). (The formation of double-stranded DNA is plotted against C0t, which is a term that combines two variables: initial DNA concentration (C0) and time of incubation (t). A solution containing a high concentration of DNA incubated for a short time will have the same C0t as one of low concentration incubated for a correspondingly longer time; both will have the same percentage of reannealed DNA.) The genome size, that is, the number of nucleotide base pairs in one copy of total genetic information of the organism, is indicated by the arrows near the upper numerical scale. The shape of each of the renaturation curves is very simple and occurs with a single slope. However, the time over which renaturation occurs is very different and depends on the concentration of complementary fragments, which in turn depends on the size of the genome. The larger the genome, the lower the concentration of complementary fragments in solution, and the greater the time required for renaturation to be complete.
908. Figure 10.17: An idealized plot showing the kinetics of renaturation of eukaryotic DNA. When single-stranded DNA is allowed to reanneal, three classes of fragments can usually be distinguished by their frequency of repetition within the genome: a highly repeated DNA fraction, a moderately repeated DNA fraction, and a nonrepeated (single-copy) DNA fraction. (Note: This is an idealized plot: the three classes of sequences are not as clearly separated in an actual renaturation curve.)
909. Highly Repeated DNA Sequences
910. Figure 10.18: DNA fingerprinting. In this technique, which is used widely to identify an individual from a sample of DNA, the DNA is digested by treatment with specific nucleases (called restriction endonucleases, described in Section 18.12), and the DNA fragments are separated on the basis of length by gel electrophoresis. The location in the gel of DNA fragments containing specific DNA sequences is determined using labeled probes with sequences complementary to those being sought. The DNA fragments that bind these probes have variable lengths from one person to the next because of the presence of variable numbers of short tandem repeats (STRs) in the genome. Forensic labs typically analyze about 13 STR markers that are known to be highly polymorphic. The chance that two individuals will have identical STR profiles is astronomically small. The fingerprint shown in this figure was used in a criminal case in which the defendant was charged with the fatal stabbing of a young woman. The bloodstains on the pants and shirt of the defendant were compared to the known blood standards from the victim and defendant. DNA from the bloodstains on the defendant’s clothing did not match his own blood standard, but they did match that of the victim. The lanes contain DNA from the following sources: 1, 2, 3, 9, and 10 are control DNA samples that serve as quality control checks; 4, the defendant’s blood; 5, bloodstains from the defendant’s pants; 6 and 7, bloodstains from the defendant’s shirt; and 8, the victim’s blood. With the advent of DNA amplification techniques (i.e., PCR), minuscule samples of DNA can be used for these analyses.
911. Figure 10.19: Fluorescence in situ hybridization and the localization of satellite DNA.
912. Moderately Repeated DNA Sequences
913. THE HUMAN PERSPECTIVE: Diseases that Result from Expansion of Trinucleotide Repeats

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